Fungi and Food Spoilage, 3rd ed

Fungi and Food Spoilage

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John I. Pitt

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Ailsa D. Hocking

Fungi and Food Spoilage

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John I. Pitt Honorary Research Fellow CSIRO Food and Nutritional Sciences North Ryde, NSW 2113 Australia [email protected]

Ailsa D. Hocking Honorary Research Fellow CSIRO Food and Nutritional Sciences North Ryde, NSW 2113 Australia [email protected]

ISBN 978-0-387-92206-5 e-ISBN 978-0-387-92207-2 DOI 10.1007/978-0-387-92207-2 Springer Dordrecht Heidelberg London New York Library of Congress Control Number: 2009920217 # Springer ScienceþBusiness Media, LLC 2009 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer ScienceþBusiness Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Springer is part of Springer ScienceþBusiness Media (www.springer.com)

Preface to the Third Edition

In contrast to the second edition, the third edition of ‘‘Fungi and Food Spoilage’’ is evolutionary rather than revolutionary. The second edition was intended to cover almost all of the species likely to be encountered in mainstream food supplies, and only a few additional species have been included in this new edition. The third edition represents primarily an updating – of taxonomy, physiology, mycotoxin production and ecology. Changes in taxonomy reflect the impact that molecular methods have had on our understanding of classification but, it must be said, have not radically altered the overall picture. The improvements in the understanding of the physiology of food spoilage fungi have been relatively small, reflecting perhaps the lack of emphasis on physiology in modern microbiological science. Much remains to be understood about the specificity of particular fungi for particular substrates, of the influence of water activity on the growth of many of the species treated, and even on such basic parameters as cardinal temperatures for growth and the influence of pH and preservatives. Since 1997, a great deal has been learnt about the specificity of mycotoxin production and in which commodities and products-specific mycotoxins are likely to occur. Changes in our understanding of the ecology of the included species are also in most cases evolutionary. A great number of papers have been published on the ecology of foodborne fungi in the past few years, but with few exceptions the basic ecology of the included species remains. Recent changes in our understanding of foodborne fungi include the realisation that Aspergillus carbonarius is a major source of ochratoxin A in the world food supply, that A. westerdijkiae and not A. ochraceus is the other common Aspergillus species making this toxin and that these species are responsible for ochratoxin A in foods outside the cool temperate regions, where Penicillium verrucosum is the important species. In recent years a number of new species have been found to be capable of producing aflatoxin, but the fact remains that most aflatoxin in the global food supply is produced by A. flavus and A. parasiticus. The taxonomy of Fusarium species is still undergoing major revision. However, the renaming of Fusarium moniliforme as F. verticillioides is the only change of importance here. Recent publications have improved our understanding of species – mycotoxin relationships within Fusarium.

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Among the colleagues who helped us to prepare this edition, we wish to particularly thank Dr Anne-Laure Markovina, now of the University of Sydney, who assisted in literature searches and some cultural and photographic work, and Mr N.J. Charley who has continued his excellent work of curating the FRR culture collection, on which so much of the descriptive work in this book is based.

Preface to the Third Edition

Preface to the First Edition

This book is designed as a laboratory guide for the food microbiologist to assist in the isolation and identification of common foodborne fungi. We emphasise the fungi which cause food spoilage, but also devote space to the fungi commonly encountered in foods at harvest, and in the food factory. As far as possible, we have kept the text simple, although the need for clarity in the descriptions has necessitated the use of some specialised mycological terms. The identification keys have been designed for use by microbiologist with little or no prior knowledge of mycology. For identification to genus level, they are based primarily on the cultural and physiological characteristics of fungi grown under a standard set of conditions. The microscopic features of the various fungi become more important when identifying isolates at the species level. Nearly all of the species treated have been illustrated with colony photographs, together with photomicrographs or line drawings. The photomicrographs were taken using a Zeiss WL microscope fitted with Nomarski interference contrast optics. We are indebted to Mr W. Rushton and Ms L. Burton, who printed the many hundreds of photographs used to make up the figures in this book. We also wish to express our appreciation to Dr D.L. Hawksworth, Dr A.H.S. Onions and Dr B.C. Sutton of the Commonwealth Mycological Institute, Kew, Surrey, UK, Professor P.E. Nelson and the staff of the Fusarium Research Center, University of Pennsylvania, USA and Dr L.W. Burgess of the University of Sydney, who generously provided facilities, cultures and advice on some of the genera studied.

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Preface to the Second Edition

In planning for the second edition of ‘‘Fungi and Food Spoilage’’, we decided that the book would benefit from a larger format, which would permit improved illustrations, and from some expansion of the text, in both numbers of species treated and overall scope. These aims have been realised. The Crown Quarto size has allowed us to include substantially larger, clearer illustrations. Many new photographs and photomicrographs have been added, the latter taken using a Zeiss Axioscop microscope fitted with Nomarski differential interference contrast optics. We have taken the opportunity to include more than 40 additional species descriptions, to add a new section on mycotoxin production for each species and to update and upgrade all of the text. Since the first edition, changes in the climate for stabilising fungal nomenclature have resulted in development of a list of ‘‘Names in Current Use’’ for some important genera, including Aspergillus and Penicillium. Names of species used in the second edition are taken from that list, which was given special status by the International Botanical Congress, Tokyo, 1994. Names used in this edition have priority over any other names for a particular species. Publication of a list of ‘‘Authors of Fungal Names’’ (P.M. Kirk and A.E. Ansell, Index of Fungi, Supplement: 1–95, 1992) has also stabilised names of authorities for all fungal species. Abbreviations of authors’ names used in this edition conform to those recommended by Kirk and Ansell. Some progress in standardisation of methods and media has also been made, primarily through the efforts of the International Commission on Food Mycology. The first edition included some 400 references. When we began revisionary work, we felt that the number of references in the area of food mycology had probably doubled or increased by perhaps 150% during the intervening years. In fact, this second edition includes over 1900 references, almost a five-fold increase over the 1985 edition! This provides a clear indication that interest in, and study of, food mycology has greatly increased in recent years. Modern referencing systems have enabled us to expand information from tropical sources, especially in Asia and Africa, but we are conscious of the fact that treatment of fungi found in foods on a worldwide basis remains rather incomplete. We gratefully acknowledge support and assistance from colleagues who have contributed to this new edition. Ms J.C. Eyles formatted and printed the camera

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ready copy, Ms C. Heenan collated, arranged and formatted the illustrations and Mr N.J. Charley looked after the culture collection, culture growth and colony photography. Without this level of support, the book would not have been completed.

Preface to the Second Edition

Contents 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

2

The Ecology of Fungal Food Spoilage . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Water Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Hydrogen Ion Concentration . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Temperature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Gas Tension . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Consistency. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Nutrient Status . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Specific Solute Effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Preservatives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Conclusions: Food Preservation . . . . . . . . . . . . . . . . . . . . . . . .

3 3 4 5 7 8 8 8 9 9

3

Naming and Classifying Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Taxonomy and Nomenclature: Biosystematics . . . . . . . . . . . . . 3.2 Hierarchical Naming . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Zygomycotina. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Ascomycotina . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Basidiomycotina . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6 The Ascomycete – Conidial Fungus Connection . . . . . . . . . . . . 3.7 Dual Nomenclature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.8 Practical Classification of Fungi . . . . . . . . . . . . . . . . . . . . . . . .

11 11 12 12 13 15 15 15 16

4

Methods for Isolation, Enumeration and Identification . . . . . . . . . . . . 4.1 Sampling. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Enumeration Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1 Direct Plating. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.2 Dilution Plating . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.3 Incubation Conditions. . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Sampling Surfaces . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Air Sampling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Isolation Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.1 Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.2 Moulds. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.3 Short Term Storage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Choosing a Suitable Medium . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6.1 General Purpose Enumeration Media. . . . . . . . . . . . . . . 4.6.2 Selective Isolation Media. . . . . . . . . . . . . . . . . . . . . . . . . 4.6.3 Techniques for Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6.4 Techniques for Heat-Resistant Fungi . . . . . . . . . . . . . . . 4.6.5 Other Plating Techniques . . . . . . . . . . . . . . . . . . . . . . . . 4.7 Estimation of Fungal Biomass . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.1 Chitin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.2 Ergosterol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.3 Impedimetry and Conductimetry . . . . . . . . . . . . . . . . . .

19 19 19 20 21 22 22 23 23 23 24 24 25 26 27 30 32 33 34 34 35 37 xi

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Contents

4.7.4 Adenosine Triphosphate (ATP) . . . . . . . . . . . . . . . . . 4.7.5 Fungal Volatiles. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.6 Immunological Techniques . . . . . . . . . . . . . . . . . . . . . 4.7.7 Molecular Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8 Identification Media and Methods. . . . . . . . . . . . . . . . . . . . . . 4.8.1 Standard Methodology . . . . . . . . . . . . . . . . . . . . . . . . 4.8.2 Plating Regimen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8.3 Inoculation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8.4 Additional Media and Methods . . . . . . . . . . . . . . . . . 4.8.5 Identification of Fusarium Species. . . . . . . . . . . . . . . . 4.8.6 Yeasts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9 Examination of Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9.1 Colony Diameters . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9.2 Colony Characters. . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9.3 Preparation of Wet Mounts for Microscopy. . . . . . . . 4.9.4 Staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9.5 Microscopes and Microscopy . . . . . . . . . . . . . . . . . . . 4.10 Preservation of Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.10.1 Lyophilisation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.10.2 Other Storage Techniques . . . . . . . . . . . . . . . . . . . . . . 4.11 Housekeeping in the Mycological Laboratory. . . . . . . . . . . . . 4.11.1 Culture Mites. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.11.2 Problem Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.11.3 Pathogens and Laboratory Safety . . . . . . . . . . . . . . . .

37 37 38 40 41 41 41 41 42 43 44 45 45 45 46 46 47 48 48 49 50 50 51 51

Primary Keys and Miscellaneous Fungi . . . . . . . . . . . . . . . . . . . . . . . . 5.1 The General Key . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.1 Notes on the General Key . . . . . . . . . . . . . . . . . . . . . . 5.2 Miscellaneous Fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Genus Acremonium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4 Genus Alternaria Nees: Fr.. . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5 Genus Arthrinium Kunze . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.6 Genus Aureobasidium Viala and G. Boyer . . . . . . . . . . . . . . . . 5.7 Genus Bipolaris Shoemaker . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.8 Genus Botrytis P. Micheli: Fr. . . . . . . . . . . . . . . . . . . . . . . . . . 5.9 Genus Chaetomium Kunze . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.10 Genus Chrysonilia Arx . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.11 Genus Cladosporium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.12 Genus Colletotrichum Corda . . . . . . . . . . . . . . . . . . . . . . . . . . 5.13 Genus Curvularia Boedijn. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.14 Genus Drechslera S. Ito . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.15 Genus Endomyces Reess . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.16 Genus Epicoccum Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.17 Genus Fusarium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.18 Genus Geotrichum Link: Fr.. . . . . . . . . . . . . . . . . . . . . . . . . . . 5.19 Genus Hyphopichia Arx and van der Walt. . . . . . . . . . . . . . . . 5.20 Genus Lasiodiplodia Ellis and Everh. . . . . . . . . . . . . . . . . . . . . 5.21 Genus Monascus Tiegh. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.22 Genus Moniliella Stolk and Dakin . . . . . . . . . . . . . . . . . . . . . . 5.23 Genus Nigrospora Zimm. . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

53 54 55 56 58 60 64 65 67 68 70 73 75 81 82 85 86 88 89 122 124 125 127 129 131

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5.24 5.25 5.26 5.27 5.28 5.29 5.30

Genus Pestalotiopsis Steyaert . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Phoma Sacc. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Stemphylium Wallr. . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Trichoconiella B.L. Jain. . . . . . . . . . . . . . . . . . . . . . . . . Genus Trichoderma Pers. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Trichothecium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Ulocladium Preuss . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

133 134 136 137 139 140 142

6

Zygomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1 Genus Absidia Tiegh. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Genus Cunninghamella Matr. . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3 Genus Mucor P. Micheli: Fr. . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4 Genus Rhizomucor (Lucet and Costantin) Vuill. . . . . . . . . . . . 6.5 Genus Rhizopus Ehrenb.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.6 Genus Syncephalastrum J. Schrot. ¨ ...................... 6.7 Genus Thamnidium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

145 148 149 151 157 158 165 167

7

Penicillium and Related Genera . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1 Genus Byssochlamys Westling . . . . . . . . . . . . . . . . . . . . . . . . . 7.2 Genus Eupenicillium F. Ludw. . . . . . . . . . . . . . . . . . . . . . . . . . 7.3 Genus Geosmithia Pitt . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4 Genus Paecilomyces Bainier . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.5 Genus Scopulariopsis Bainier . . . . . . . . . . . . . . . . . . . . . . . . . . 7.6 Genus Talaromyces C.R. Benj.. . . . . . . . . . . . . . . . . . . . . . . . . 7.7 Genus Penicillium Link. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.7.1 Penicillium subgenus Aspergilloides Dierckx . . . . . . . . . 7.7.2 Penicillium subgenus Furcatum Pitt. . . . . . . . . . . . . . . . 7.7.3 Penicillium subgenus Penicillium . . . . . . . . . . . . . . . . . . 7.7.4 Penicillium subgenus Biverticillium Dierckx . . . . . . . . .

169 170 175 182 183 187 188 194 196 207 223 263

8

Aspergillus and Related Teleomorphs. . . . . . . . . . . . . . . . . . . . . . . . . . 8.1 Genus Emericella Berk. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2 Genus Eurotium Link: Fr. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3 Genus Neosartorya Malloch and Cain . . . . . . . . . . . . . . . . . . . 8.4 Genus Aspergillus Fr.: Fr. . . . . . . . . . . . . . . . . . . . . . . . . . . . .

275 279 281 292 295

9

Xerophiles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.1 Genus Basipetospora G.T. Cole and W.B. Kendr. . . . . . . . . . . 9.2 Genus Chrysosporium Corda . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3 Genus Eremascus Eidam . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4 Genus Polypaecilum G. Sm. . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.5 Genus Wallemia Johan-Olsen. . . . . . . . . . . . . . . . . . . . . . . . . . 9.6 Genus Xeromyces L.R. Fraser . . . . . . . . . . . . . . . . . . . . . . . . .

339 340 342 347 348 350 353

10

Yeasts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

357

11

Fresh and Perishable Foods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.1 Spoilage of Living, Fresh Foods . . . . . . . . . . . . . . . . . . . . . . . 11.2 Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.1 Citrus Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.2 Pome Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.3 Stone Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

383 383 383 384 385 386

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11.3

11.4 11.5 11.6

12

11.2.4 Tomatoes and other Solanaceous Fruit. . . . . . . . . . . 11.2.5 Melons and other Cucurbits . . . . . . . . . . . . . . . . . . . 11.2.6 Grapes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.7 Berries . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.8 Figs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.9 Tropical Fruit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vegetables . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.1 Peas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.2 Beans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.3 Onions and Garlic . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.4 Potatoes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.5 Roots and Tubers . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.6 Yams . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.7 Cassava . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.8 Leafy and other Green Vegetables . . . . . . . . . . . . . . Dairy Foods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Meats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cereals, Nuts and Oilseeds . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.1 Wheat, Barley and Oats. . . . . . . . . . . . . . . . . . . . . . . 11.6.2 Rice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.3 Maize . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.4 Soybeans and Mung Beans . . . . . . . . . . . . . . . . . . . . 11.6.5 Other Beans and Pulses . . . . . . . . . . . . . . . . . . . . . . . 11.6.6 Sunflower Seed . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.7 Sorghum. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.8 Peanuts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.9 Cashews and Brazil Nuts . . . . . . . . . . . . . . . . . . . . . . 11.6.10 Almonds, Hazelnuts, Walnuts and Pecans . . . . . . . . 11.6.11 Pistachios . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.12 Copra . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Spoilage of Stored, Processed and Preserved Foods . . . . . . . . . . . . . . . 12.1 Low Water Activity Foods: Dried Foods . . . . . . . . . . . . . . . . 12.1.1 Cereals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.2 Flour . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.3 Pasta. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.4 Bakery Products. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.5 Maize . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.6 Soybeans, Mung Beans, other Beans and Chickpeas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.7 Nuts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.8 Peanuts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.9 Hazelnuts, Walnuts, Pecans and Almonds . . . . . . . . 12.1.10 Pistachio Nuts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.11 Other Nuts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.12 Coconut . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.13 Spices . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.14 Coffee Beans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.15 Cocoa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.16 Dried Meat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

387 387 388 389 389 390 391 391 391 391 392 392 392 392 393 393 394 395 395 396 396 397 398 398 398 398 399 399 399 400 401 401 402 403 403 403 404 405 406 406 407 408 409 409 410 410 411 412

Contents

xv

12.2

Low Water Activity Foods: Concentrated Foods . . . . . . . . . . 12.2.1 Jams . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.2 Dried Fruit. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.3 Fruit Cakes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.4 Confectionery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.5 Fruit Concentrates. . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.6 Honey and Syrups . . . . . . . . . . . . . . . . . . . . . . . . . . . . Low Water Activity Foods: Salt Foods . . . . . . . . . . . . . . . . . . Intermediate Moisture Foods: Processed Meats . . . . . . . . . . . Heat Processed Acid Foods . . . . . . . . . . . . . . . . . . . . . . . . . . . Preserved Foods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cheese . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

412 412 413 414 415 415 416 416 417 418 418 419

Media Appendix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

423

Glossary. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

427

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

431

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

503

12.3 12.4 12.5 12.6 12.7

Chapter 1

Introduction

From the time when primitive man began to cultivate crops and store food, spoilage fungi have demanded their tithe. Fuzzes, powders and slimes of white or black, green, orange, red and brown have silently invaded – acidifying, fermenting, discolouring and disintegrating, rendering nutritious commodities unpalatable or unsafe. Until recently, fungi have generally been regarded as causing only unaesthetic spoilage of food, despite the fact that Claviceps purpurea was linked to human disease more than 200 years ago, and the acute toxicity of macrofungi has long been known. Japanese scientists recognised the toxic nature of yellow rice 100 years ago, but 70 years elapsed before its fungal cause was confirmed. Alimentary toxic aleukia killed many thousands of people in the USSR in 1944–1947; although fungal toxicity was suspected by 1950, the causal agent, T-2 toxin, was not clearly recognised for another 25 years. Forgacs and Carll (1952), in a prophetic article, warned of the danger from common spoilage fungi, but it was not until 1960, when the famous "Turkey X" disease killed 100,000 turkey poults in Great Britain, and various other disasters followed in rapid succession, that the Western world became aware that common spoilage moulds could produce significant toxins. Since 1960 a seemingly endless stream of toxigenic fungi and potentially toxic compounds has been discovered. On these grounds alone, the statement ‘‘It’s only a mould’’ is no longer acceptable to food microbiologist, health inspector or consumer. The demand for accurate identification and characterisation of food spoilage fungi has become urgent. In the flurry of research into mycotoxins, however, it must not be forgotten that food spoilage as such

remains an enormous problem throughout the world. Figures are difficult to obtain. However, even given a dry climate and advanced technology, losses of food to fungal spoilage in Australia must be in excess of $10,000,000 per annum: losses in humid tropical climates and countries with less highly developed technology remain staggering. An estimate of 5–10% of all food production is not unrealistic. Research into fungal food spoilage and its prevention is clearly an urgent necessity: lacking in spectacular appeal, it is, however, often neglected. A further point, of the highest significance, needs emphasis here. Research on the fungi which cause food spoilage, and the mycotoxins they produce, can only be carried out effectively if based on accurate identification of the microorganisms responsible. Taxonomy and nomenclature (systematics) make up the vital root system of all the trees of biological science. The prevention of fungal food spoilage as an art is old, but as a discipline, young. Drying, the oldest method of food preservation, has been practiced for millennia and is still the most common, effective and cheap technique for preserving food. Only recently have we been able to identify with certainty the fungal species which cause spoilage of dried foods. Prediction of their responses to a given environment, specified by physico-chemical parameters such as water activity, temperature, pH and oxygen tension, even now is often uncertain. Within historic times, newer methods of food preservation have been introduced – salting, curing, canning, refrigeration, freezing, the use of preservatives, irradiation and most recently, high hydrostatic pressure. Freezing excepted, each new technique has selected for one or more fungal species resistant to

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_1, Ó Springer ScienceþBusiness Media, LLC 2009

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the process applied. As examples we can take Polypaecilum pisce on salt fish, Xeromyces bisporus on fruit cake, Cladosporium herbarum on refrigerated meat, Zygosaccharomyces bailii in preserved juices, Z. rouxii in jams and fruit concentrates, Aspergillus flavus on peanuts, Eurotium chevalieri on hazel nuts, Penicillium roqueforti on cheeses, Byssochlamys fulva in acid canned foods . . . the list of quite specific food – fungus associations is extensive. The study of such associations is one of the more important branches of the young discipline, food mycology. This book sets out to document current knowledge on the interaction of foods and fungi, in the context of spoilage and toxicity, not food production or biotechnology. Four aspects are examined. First, ecology: what factors in foods select for particular kinds of fungi? A chapter is devoted to the physical and chemical parameters which influence the growth of fungi in foods. Second, methodology: how do we isolated fungi from foods? What are the best media to use? How do we go about identifying food spoilage fungi? Third, the commodity: what fungi are usually associated with a particular food? Here ecological factors interact to produce a more or less specific habitat. Major classes of foods and their associated spoilage fungi are described. Finally, the fungus: what fungus is that? In a series of chapters, the main food spoilage moulds and yeasts are described and keyed, together with others commonly associated with food but not noted for spoilage. Where possible, further information is

1

Introduction

given on known habitats and sources, physiology, heat resistance, etc., together with a selective bibliography. Accurate information on mycotoxin production is also included. As far as possible, the precise terminology for fungal structures used by the pure mycologist and indeed most necessary for him has been avoided in these chapters. Some concepts and terms are of course essential: these have been introduced as needed and are listed in a glossary. The taxonomic sections of this book are designed to facilitate identification of food spoilage and common food contaminant fungi. A standardised plating regimen is used, originally developed for the identification of Penicillium species (Pitt, 1979b) and extended here to other genera relevant to the food industry. Under this regimen, cultures are incubated for 1 week at 5, 25 and 378C on a single standard medium and at 258C on two others. In conjunction with the appropriate keys, this system will enable identification of most foodborne fungi to species level in just 7 days. For a few kinds of fungi, notably yeasts and xerophiles, subsequent growth under other more specialised conditions will be necessary. Finally, this book is dedicated to the general food microbiologist. May it help to restore equilibrium and assist in continued employment, when the quality assurance manager demands: ‘‘What is it?’’ . . . ‘‘How did it get in?’’ . . . ‘‘What does it do?’’ . . . ‘‘How do we get rid of it?’’ . . . and, worst of all . . . ‘‘Is it toxic?’’

Chapter 2

The Ecology of Fungal Food Spoilage

Food is not commonly regarded as an ecosystem, perhaps on the basis that it is not a ‘‘natural’’ system. Nevertheless an ecosystem it is and an important one, because food plants and the fungi that colonise their fruiting parts (seeds and fruit) have been co-evolving for millennia. The seed and nut caches of rodents have provided a niche for the development of storage fungi. Fallen fruit, as they go through the cycle of decay and desiccation, have provided substrate for a range of fungi. Humans have aided and abetted the development of food spoilage fungi through their vast and varied food stores. It can be argued, indeed, some rapidly evolving organisms, such as haploid asexual fungi, are moving into niches created by man’s exploitation of certain plants as food. Food by its very nature is expected to be nutritious: therefore, food is a rich habitat for microorganisms, in contrast with the great natural systems, soil, water and plants. Given the right physico-chemical conditions, only the most fastidious microorganisms are incapable of growth in foods, so that factors other than nutrients usually select for particular types of microbial populations. Perhaps the most important of these factors relates to the biological state of the food. Living foods, particularly fresh fruits, vegetables, and also grains and nuts before harvest, possess powerful defence mechanisms against microbial invasion. The study of the spoilage of such fresh foods is more properly a branch of plant pathology than food microbiology. The overriding factor determining spoilage of a fresh, living food is the ability of specific microorganisms to overcome defence mechanisms. Generally speaking, then, spoilage of

fresh foods is limited to particular species. Such specific relationships between fresh food and fungus are discussed in Chapter 11 and under particular species. Other kinds of foods are moribund, dormant or nonliving, and the factors which govern spoilage are physical and chemical. There are eight principal factors: (l) water activity; (2) hydrogen ion concentration; (3) temperature – of both processing and storage; (4) gas tension, specifically of oxygen and carbon dioxide; (5) consistency; (6) nutrient status; (7) specific solute effects; and (8) preservatives. Each will be discussed in turn below.

2.1 Water Activity Water availability in foods is most readily measured as water activity. Water activity (aw), is a physicochemical concept, introduced to microbiologists by Scott (1957), who showed that aw effectively quantified the relationship between moisture in foods and the ability of microorganisms to grow on them. Water activity is defined as a ratio: aw ¼ p=po ; where p is the partial pressure of water vapour in the test material and po is the saturation vapour pressure of pure water under the same conditions.

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_2, Ó Springer ScienceþBusiness Media, LLC 2009

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Water activity is numerically equal to equilibrium relative humidity (ERH) expressed as a decimal. If a sample of food is held at constant temperature in a sealed enclosure until the water in the sample equilibrates with the water vapour in the enclosed air space (Fig. 2.1a), then aw ðfoodÞ ¼ ERH ðairÞ=100: Conversely, if the ERH of the air is controlled in a suitable way, as by a saturated salt solution, at equilibrium the aw of the food will be numerically equal to the generated ERH (Fig. 2.1b). In this way, aw can be experimentally controlled, and the relation of aw to moisture (the sorption isotherm) can be studied. For further information on water activity, its measurement and significance in foods see Duckworth (1975); Pitt (1975); Troller and Christian (1978); Rockland and Beuchat (1987). In many practical situations, aw is the dominant environmental factor governing food stability or spoilage. A knowledge of fungal water relations will then enable prediction both of the shelf life of foods and of potential spoilage fungi. Although the water relations of many fungi will be considered individually in later chapters, it is pertinent here to provide an overview. Like all other organisms, fungi are profoundly affected by the availability of water. On the aw scale, life as we know it exists over the range 0.9999þ to 0.60 (Table 2.1). Growth of animals is virtually confined to 1.0–0.99 aw; the permanent wilt point of mesophytic plants is near 0.98 aw; and most microorganisms cannot

Fig. 2.1 The concept of water activity (aw) (a) the relationship between aw and equilibrium relative humidity (ERH); (b) one method of controlling aw by means of a saturated salt solution, which generates a specific ERH at a specific constant temperature

The Ecology of Fungal Food Spoilage

grow below 0.95 aw. A few halophilic algae and bacteria can grow in saturated sodium chloride (0.75 aw), but are confined to salty environments. Ascomycetous fungi and conidial fungi of ascomycetous origin comprise most of the organisms capable of growth below 0.9 aw. Fungi capable of growth at low aw, in the presence of extraordinarily high solute concentrations both inside and out, must be ranked as among the most highly evolved organisms on earth. Even among the fungi, this evolutionary path must have been of the utmost complexity: the ability to grow at low aw is confined to only a handful of genera (Pitt, 1975). The degree of tolerance to low aw is most simply expressed in terms of the minimum aw at which germination and growth can occur. Fungi able to grow at low aw are termed xerophiles: one widely used definition is that a xerophile is a fungus able to grow below 0.85 aw under at least one set of environmental conditions (Pitt, 1975). Xerophilic fungi will be discussed in detail in Chapter 9. Information about the water relations of many fungi remains fragmentary, but where it is known it has been included in later chapters.

2.2 Hydrogen Ion Concentration At high water activities, fungi compete with bacteria as food spoilers. Here pH plays the decisive role. Bacteria flourish near neutral pH and fungi cannot compete unless some other factor, such as low water

2.3 Temperature

5

Table 2.1 Water activity and microbial water relations in perspectivea aw

Perspective

Foods

1.00

Blood, plant wilt point, seawater

0.95

Most bacteria

Vegetables meat, milk fruit Bread

0.90 0.85 0.80

Staphylococcus aureus

Ham Dry salami

0.75

Salt lake Halophiles

0.70

Jams Salt fish Fruit cake Confectionery Dried fruit Dry grains

Moulds

Yeasts

Basidiomycetes Most soil fungi Mucorales Fusarium Rhizopus, Cladosporium Aspergillus flavus Xerophilic Penicillia Xerophilic Aspergilli Wallemia Eurotium Chrysosporium Eurotium halophilicum

Basidiomycetes Most ascomycetes Zygosaccharomyces rouxii (salt) Zygosaccharomyces bailii Debaryomyces hansenii

0.65 Xeromyces bisporus Zygosaccharomyces rouxii (sugar) 0.60 DNA disordered a Modified from data of J.I. Pitt as reported by Brown (1974). Water activities shown for microorganisms approximate minima for growth reported in the literature.

activity or a preservative, renders the environment hostile to the bacteria. As pH is reduced below about 5, growth of bacteria becomes progressively less likely. Lactic acid bacteria are exceptional, as they remain competitive with fungi in some foods down to about pH 3.5. Most fungi are little affected by pH over a broad range, commonly 3–8 (Wheeler et al., 1991). Some conidial fungi are capable of growth down to pH 2, and yeasts down to pH 1.5. However, as pH moves away from the optimum, usually about pH 5, the effect of other growth limiting factors may become apparent when superimposed on pH. Figure 2.2 is an impression of the combined influence of pH and aw on microbial growth: few accurate data points exist and the diagram is schematic. For heat-processed foods, pH 4.5 is of course critical: heat processing to destroy the spores of Clostridium botulinum also destroys all fungal spores. In acid packs, below pH 4.5, less severe processes may permit survival of heat-resistant fungal spores (Section 2.3).

2.3 Temperature The influence of temperature in food preservation and spoilage has two separate facets: temperatures during processing and those existing during storage.

As noted above, heat-resistant fungal spores may survive pasteurising processes given to acid foods. Apart from a few important species, little information exists on the heat resistance of fungi. Much of the information that does exist must be interpreted with care, as heating menstrua and conditions can vary markedly, and these may profoundly affect heat resistance. High levels of sugars are generally protective (Beuchat and Toledo, 1977). Low pH and preservatives increase the effect of heat (Beuchat, 1981a, b; Rajashekhara et al., 2000) and also hinder resuscitation of damaged cells (Beuchat and Jones, 1978). Ascospores of filamentous fungi are more heat resistant than conidia (Pitt and Christian, 1970; Table 2.2). Although not strictly comparable, data of Put et al. (1976) indicate that the heat resistance of yeast ascospores and vegetative cells is of the same order as that of fungal conidia. Among the ascomycetous fungi, Byssochlamys species are notorious for spoiling heat processed fruit products (Olliver and Rendle, 1934; Richardson, 1965). The heat resistance of B. fulva ascospores varies markedly with isolate and heating conditions (Beuchat and Rice, 1979): a D value between 1 and 12 min at 908C (Bayne and Michener, 1979) and a z value of 6–7C8 (King et al., 1969) are practical working figures. The heat resistance of

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2

The Ecology of Fungal Food Spoilage

Fig. 2.2 A schematic diagram showing the combined influence of water activity and pH on microbial growth

B. nivea ascospores is marginally lower (Beuchat and Rice, 1979; Kotzekidou, 1997a). Ascospores of Neosartorya fischeri have a similar heat resistance to those of Byssochlamys fulva, but have been reported less frequently as a cause of food spoilage. Heat resistant fungi are discussed further in Chapter 4. Food products may be stored at ambient temperatures, in which case prevention of spoilage relies on other parameters, or under refrigeration, where temperature is expected to play a preservative

role. Food frozen to –108C or below appears to be microbiologically stable, despite some reports of fungal growth at lower temperatures. The lowest temperatures for fungal growth are in the range –7 to 08C, for species of Fusarium, Cladosporium, Penicillium and Thamnidium (Pitt and Hocking, 1997). Nonsterile food stored at ca. 58C in domestic refrigerators, where conditions of high humidity prevail, will eventually be spoiled by fungi of these genera. At high aw and neutral pH, psychrophilic bacteria may also be important (mostly Pseudomonas species).

Table 2.2 Comparative heat resistance of ascospores and conidiaa Survivors (%) Fungus

Spore type

Initial viable count/ml

Ascospores 5.0  102 Conidia 7.3  102 Eurotium chevalieri Ascospores 1.0  103 Conidia 8.9  102 Xeromyces bisporus Ascospores 1.0  103 Aspergillus candidus Conidia 3.8  102 Wallemia sebi Conidia 7.1  102 a Heated at temperatures shown for 10 min. Data from Pitt and Christian (1970). Eurotium amstelodami

508C

608C

708C

93 107 103 128 93 102 42

85 0.3 62 0.1 30 0 0

3 0 21 0 0.3 0 0

2.4 Gas Tension

Thermophilic fungi, i.e. those which grow only at high temperatures, are rarely of significance in food spoilage. If overheating of commodities occurs, however, in situations such as damp grain, thermophiles can be a very serious problem. Thermotolerant fungi, i.e. species able to grow at both moderate and high temperatures, are of much greater significance. Aspergillus flavus and A. niger, able to grow between ca. 8 and 458C, are among the most destructive moulds known.

2.4 Gas Tension Food spoilage moulds, like almost all other filamentous fungi, have an absolute requirement for oxygen. However, many species appear to be efficient oxygen scavengers, so that the total amount of oxygen available, rather than the oxygen tension, determines growth. The concentration of oxygen dissolved in the substrate has a much greater influence on fungal growth than atmospheric oxygen tension (Miller and Golding, 1949). For example, Penicillium expansum grows virtually normally in 2.1% oxygen over its entire temperature range (Golding, 1945), and many other common food spoilage fungi are inhibited only slightly when grown in nitrogen atmospheres containing approximately 1.0% oxygen (Hocking, 1990). Paecilomyces variotii produced normal colonies at 258C under 650 mm of vacuum (Pitt, unpublished). Most food spoilage moulds appear to be sensitive to high levels of carbon dioxide, although there are notable exceptions. When maintained in an atmosphere of 80% carbon dioxide and 4.2% oxygen, Penicillium roqueforti still grew at 30% of the rate in air (Golding, 1945), provided that the temperature was above 208C. In 40% CO2 and 1% O2, P. roqueforti grew at almost 90% of the rate in air (Taniwaki et al., 2001a). Xeromyces bisporus has been reported to grow in similar levels of carbon dioxide (Dallyn and Everton, 1969). Byssochlamys species appear to be particularly tolerant of conditions of reduced oxygen and/or elevated carbon dioxide. Growth of Byssochlamys nivea was little affected by replacement of nitrogen in air by carbon dioxide, and growth in carbon dioxide–air mixtures was proportional only to

7

oxygen concentration, at least up to 90% carbon dioxide (Yates et al., 1967). Both Byssochlamys nivea and B. fulva were capable of growth in atmospheres containing 20, 40 or 60% carbon dioxide with less than 0.5% oxygen, but inhibition increased with increasing carbon dioxide concentration (Taniwaki et al., 2001a). Byssochlamys fulva is capable of growth in 0.27% oxygen, but not in its total absence (King et al., 1969). It is also capable of fermentation in fruit products, but presumably only if some oxygen is present. At least some species of Mucor, Rhizopus and Fusarium are able to grow and ferment in bottled liquid products and sometimes cause fermentative spoilage. Growth under these conditions may be yeast-like. Species of Mucor, Rhizopus and Amylomyces used as starter cultures in Asian fermented foods can grow under anaerobic conditions, demonstrated by growth in an anaerobe jar with a hydrogen and carbon dioxide generator (Hesseltine et al., 1985). Other authors have reported growth under anaerobic conditions of such fungi as Mucor species, Absidia spinosa, Geotrichum candidum, Fusarium oxysporum and F. solani (Stotzky and Goos, 1965; Curtis, 1969; Taniwaki, 1995). The yeast-like fungus Moniliella acetoabutans can cause fermentative spoilage under totally anaerobic conditions (Stolk and Dakin, 1966). As a generalisation, however, it is still correct to state that most food spoilage problems due to filamentous fungi occur under aerobic conditions, or at least where oxygen tension is appreciable, due to leakage or diffusion through packaging. In contrast, Saccharomyces species, Zygosaccharomyces species and other fermentative yeasts are capable of growth in the complete absence of oxygen. Indeed, S. cerevisiae and Z. bailii can continue fermentation under several atmospheres pressure of carbon dioxide. This property of S. cerevisiae has been harnessed by mankind for his own purposes, in the manufacture of bread and many kinds of fermented beverages. Z. bailii, on the other hand, is notorious for its ability to continue fermenting at reduced water activities in the presence of high levels of preservatives. Fermentation of juices and fruit concentrates may continue until carbon dioxide pressure causes container distortion or explosion. The closely related species Zygosaccharomyces rouxii is a xerophile and causes

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spoilage of low-moisture liquid or packaged products such as fruit concentrates, jams and dried fruit. The difference in oxygen requirements between moulds and fermentative yeasts is one of the main factors determining the kind of spoilage a particular commodity will undergo.

2.5 Consistency Consistency, like gas tension, exerts considerable influence over the kind of spoilage to which a food is susceptible. Generally speaking, yeasts cause more obvious spoilage in liquid products, because single celled microorganisms are able to disperse more readily in liquids. Moreover, a liquid substrate is more likely to give rise to anaerobic conditions and fermentation is more readily seen in liquids. In contrast, filamentous fungi are assisted by a firm substrate, and ready access to oxygen. The foregoing is not intended to suggest that yeasts cannot spoil solid products nor moulds liquids: merely that all other factors being equal, fermentative yeasts have a competitive advantage in liquids and cause more obvious spoilage under these conditions.

2.6 Nutrient Status As noted in the preamble to this chapter, the nutrient status of most foods is adequate for the growth of any spoilage microorganism. Generally speaking, however, it appears that fungal metabolism is best suited to substrates high in carbohydrates, whereas bacteria are more likely to spoil proteinaceous foods. Lactobacilli are an exception. Most common mould species appear to be able to assimilate any food-derived carbon source with the exception of hydrocarbons and highly condensed polymers such as cellulose and lignin. Most moulds are equally indifferent to nitrogen source, using nitrate, ammonium ions or organic nitrogen sources with equal ease. Some species achieve only limited growth if amino acids or proteins must provide both carbon and nitrogen. A few isolates classified in Penicillium subgen. Biverticillium are unable to utilise nitrate (Pitt, 1979b).

2

The Ecology of Fungal Food Spoilage

Some xerophilic fungi are known to be more demanding. Ormerod (1967) showed that growth of Wallemia sebi was strongly stimulated by proline. Xerophilic Chrysosporium species and Xeromyces bisporus also require complex nutrients, but the factors involved have not been defined (Pitt, 1975). Yeasts are often fastidious. Many are unable to assimilate nitrate or complex carbohydrates; a few, Zygosaccharomyces bailii being an example, cannot grow with sucrose as a sole source of carbon. Some require vitamins. These factors limit to some extent the kinds of foods susceptible to spoilage by yeasts. A further point on nutrients in foods is worth making here. Certain foods (or nonfoods) lack nutrients essential for the growth of spoilage fungi. Addition of nutrient, for whatever reason, can turn a safe product into a costly failure. Two cases from our own experience illustrate this point, both involving spoilage by the preservativeresistant yeast Zygosaccharomyces bailii. In the first, a highly acceptable (and nutritious) carbonated beverage containing 25% fruit juice was eventually forced from the Australian market because it was impractical to prepare it free of occasional Z. bailii cells. Effective levels of preservative could not be added legally and pasteurisation damaged its flavour. Substitution of the fruit juice with artificial flavour and colour removed the nitrogen source for the yeast. A spoilage free product resulted, at the cost of any nutritional value and a great reduction in consumer acceptance. The other case concerned a popular water-ice confection, designed for home freezing. This confection contained sucrose as a sweetener and a preservative effective against yeasts utilising sucrose. One production season the manufacturer decided, for consumer appeal, to add glucose to the formulation. The glucose provided a carbon source for Zygosaccharomyces bailii, and as a result several months production, valued at hundreds of thousands of dollars, was lost due to fermentative spoilage.

2.7 Specific Solute Effects As stated earlier, microbial growth under conditions of reduced water availability is most satisfactorily described in terms of aw. However,

2.9 Conclusions: Food Preservation

the particular solutes present in foods can exert additional effects on the growth of fungi. Scott (1957) reported that Eurotium (Aspergillus) amstelodami grew 50% faster at its optimal aw (0.96) when aw was controlled by glucose rather than magnesium chloride, sodium chloride or glycerol. Pitt and Hocking (1977) showed a similar effect for Eurotium chevalieri and reported that the extreme xerophiles Chrysosporium fastidium and Xeromyces bisporus grew poorly if at all in media containing sodium chloride as the major solute. In contrast Pitt and Hocking (1977) and Hocking and Pitt (1979) showed that germination and growth of several species of Aspergillus and Penicillium was little affected when medium aw was controlled with glucose–fructose, glycerol or sodium chloride. Zygosaccharomyces rouxii, the second most xerophilic organism known, has been reported to grow down to 0.62 aw in fructose (von Schelhorn, 1950). Its minimum aw for growth in sodium chloride is reportedly much higher, 0.85 aw (Onishi, 1963). Some fungi are halophilic, being well adapted to salty environments such as salted fish. Basipetospora halophila and Polypaecilum pisce grow more rapidly in media containing NaCl as controlling solute (Andrews and Pitt, 1987; Wheeler et al., 1988c). Such fungi have been called halophilic xerophiles to distinguish them from obligately halophilic bacteria.

2.8 Preservatives Obviously, preservatives for use in foods must be safe for human consumption. Under this constraint, food technologists in most countries are limited to the use of weak acid preservatives: benzoic, sorbic, nitrous, sulphurous, acetic and propionic acids – or, less commonly, their esters. In the concentrations permitted by most food laws, these acids are useful only at pH levels up to their pKa plus one pH unit, because to be effective they must be present as the undissociated acid. For studies of the mechanism of action of weak acid preservatives see Warth (1977, 1991); Brul and Coote (1999); Stratford and Anslow (1998) and Stratford and Lambert (1999).

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The use of chemical preservatives in foods is limited by law in most countries to relatively low levels and to specific foods. A few fungal species possess mechanisms of resistance to weak acid preservatives, the most notable being Zygosaccharomyces bailii. This yeast is capable of growth and fermentation in fruit-based cordials of pH 2.9–3, of 458C Brix and containing 800 mg/L of benzoic acid (Pitt and Hocking, 1997). The yeast-like fungus Moniliella acetoabutans can grow in the presence of 4% acetic acid and survive in 10% (Pitt and Hocking, 1997). Of the filamentous fungi, Penicillium roqueforti appears to be especially resistant to weak acid preservatives and this property has been suggested as a useful aid to isolation and identification (Engel and Teuber, 1978).

2.9 Conclusions: Food Preservation It is evident from the above discussion that the growth of fungi in a particular food is governed largely by a series of physical and chemical parameters, and definition of these can assist greatly in assessing the food’s stability. The situation in practice is made more complex by the fact that such factors frequently do not act independently, but synergistically. If two or more of the factors outlined above act simultaneously, the food may be safer than expected. This has been described by Leistner and Rodel (1976) as the ‘‘hurdle concept’’. ¨ This concept has been evaluated carefully for some commodities such as fermented sausages and is now widely exploited in the production of shelf stable bakery goods and acid sauces. For most fungi, knowledge remains meagre about the influence of the eight parameters discussed here on germination and growth. However, sufficient information is now available that some rationale for spoilage of specific commodities by certain fungi can be attempted, especially where one or two parameters are of overriding importance. This topic is considered in later chapters devoted to particular commodities.

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Chapter 3

Naming and Classifying Fungi

As with other living organisms, the name applied to any fungus is a binomial, a capitalised genus name followed by a lower case species name, both written in italics or underlined. The classification of organisms in genera and species was a concept introduced by Linneaus in 1753 and it is the keystone of biological science. It is as fundamental to the biologist as Arabic decimal numeration is to the mathematician. Here the analogy ends: the concept of ‘‘base 10’’ is rigorous; the concept of a species, fundamental as it is, is subjective and dependent on the knowledge and concepts of the biologist who described it.

3.1 Taxonomy and Nomenclature: Biosystematics Once biologists began to describe species and to assemble them into genera, questions about their relationships began to arise: is species x described by Jones in 1883 the same as species y described by Smith in 1942? Does species z, clearly distinct from x and y in some characters, belong to the same genus? The study of these relationships is termed taxonomy. Modern taxonomy is based on sound scientific principles, but still involves subjective judgment. When the decision is made that species x and species y are the same, however, the taxonomist must follow clearly established procedures in deciding which name must be used (‘‘has priority’’). The application of these procedures is termed nomenclature and, for fungi, plants and algae, is governed by the International Code of Botanical Nomenclature (ICBN). The ICBN is a relatively complex document of about 70 Articles dealing with all aspects of

correctly naming plants, algae and fungi. It is amended every 6 years by special sessions at each International Botanical Congress and is republished thereafter. The 17th version of the ICBN (the Vienna code) is the most recently published (McNeill et al., 2006). The ICBN impinges only indirectly on the work of the practicing mycologist or microbiologist. It is nevertheless of vital importance to the orderly naming of all plant life; to ignore the ICBN is to invite chaos. Where confusion arises over the correct name for a botanical species – a constant source of irritation to the nontaxonomist – it stems usually from one of three causes: indecision by, or disagreement among, taxonomists on what constitutes a particular species; incorrect application of the provisions of the ICBN; or ignorance of earlier literature. To return to our example, when species x and species y are seen to be the same, x has priority because it was published earlier; y becomes a synonym of x. Important synonyms are often listed after a name to aid the user of a taxonomy, and this procedure has been followed here. Through ignorance, the same species name may be used more than once, for example, Penicillium thomii Maire 1915 and P. thomii K.M. Zalessky 1927. The name P. thomii has been given to two quite different fungi. Clearly P. thomii Maire has priority; the later name is not valid. To avoid ambiguity, correct practice in scientific publication is to cite the author of a species at first mention, and before any formal description. The ICBN provides rules to govern change of genus name also. In our example, if species z is transferred to the genus to which species x and y belong, it retains its species name but takes the new

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_3, Ó Springer ScienceþBusiness Media, LLC 2009

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Naming and Classifying Fungi

genus name. The original author of the name z is placed in brackets after the species name, followed by the name of the author who transferred it to the correct genus. For example, Citromyces glaber Wehmer 1893 became Penicillium glabrum (Wehmer) Westling 1893 on transfer to Penicillium by Westling in 1911. Note the use of Latinised names: glaber (masculine) became glabrum (neuter) to agree with the gender of the genus to which it was transferred. Further points on the use of the ICBN arise from this example. P. glabrum retains its date of original publication, and therefore takes priority over P. frequentans Westling 1911 if the two species are combined. When Raper and Thom (1949) combined the two species, a taxonomically correct decision, they retained the name P. frequentans, which was nomenclaturally incorrect, causing confusion when Subramanian (1971) and Pitt (1979b) took up the correct name. It is worth pointing out that the confusion in this and similar situations arose from Raper and Thom’s action in ignoring the provisions of the ICBN, not from that of later taxonomists who correctly interpreted it.

connection between the Ascomycetes, the fungi that produce sexual spores in sacks, and the Deuteromycetes, where spores are always asexual, has been known for a long time. However, molecular taxonomy has provided the fundamental assurance needed to make this change. From the point of view of the food mycologist, this is a mixed blessing. The demands of the molecular systematists may yet make the taxonomy of foodborne fungi even more complicated. The taxonomic system used here is believed to be both practical and in line with the current ‘‘best practice’’ of the nomenclaturalists. The hierarchical subdivisions in Kingdom Fungi of interest in the present context are shown below, using as examples three genera and species important in food spoilage:

3.2 Hierarchical Naming

Note that names of genera, species and varieties are italicised or underlined, while higher taxonomic ranks are not. Three subkingdoms of the kingdom Fungi include genera of significance in food spoilage. As indicated in the examples above, these are Zygomycotina, Ascomycotina and (much less commonly) Basidiomycotina. Fungi from each of these subkingdoms have quite distinct properties, shared with other genera and species from the same subkingdom. Unlike other texts, this book will not rely on initial recognition of a correct subkingdom before identification of genus and species can be undertaken. Nevertheless, identification of the subkingdom can provide valuable information about a fungus, so the principal properties of these three subkingdoms are described below.

A given biological entity, or taxon in modern terminology, can be given a hierarchy of names: a cluster of related species is grouped in a genus, of related genera in families, of families in orders, orders in classes, and classes in subkingdoms. Similarly a species can be divided into smaller entities: subspecies, varieties and formae speciales (a term usually reserved for plant pathogens). In most modern classifications, the fungi are ranked, like plants and animals, as a separate kingdom. Traditionally, fungi have been divided into several subkingdoms, based on spore type and some environmental considerations. Modern molecular methods have revolutionised this. Fungi have been shown to be more closely related to animals than plants, where traditional taxonomy has always placed them. Some of the so-called ‘‘lower fungi’’ have been shown not to be fungi of all (though mycologists will no doubt continue to study them). The most important change from the point of view of the food mycologist is the demise of the subkingdom Deuteromycotina, and its absorption (almost entirely) into the Ascomycotina. The

Kingdom Subkingdom Class Order Family Genus Species Variety

Fungi Zygomycotina Zygomycetes Mucorales Mucoraceae Rhizopus stolonifer

Fungi Ascomycotina Plectomycetes Eurotiales Trichocomaceae Eurotium chevalieri intermedius

Fungi Basidiomycotina Wallemiomycetes Wallemiales Wallemiaceae Wallemia sebi

3.3 Zygomycotina Most fungi within the subkingdom Zygomycotina belong to the class Zygomycetes. Fungi in this class possess three distinctive properties:

3.4 Ascomycotina

1. Rapid growth. Most isolates grow very rapidly, often filling a Petri dish of malt extract agar with loose mycelium in 2–4 days. 2. Nonseptate mycelium. Actively growing mycelia are without septa (cross walls) and are essentially unobstructed. This allows rapid movement of cell contents, termed ‘‘protoplasmic streaming’’, which can be seen readily by transmitted light under the binocular microscope. In wet mounts the absence of septa is usually obvious (Fig. 3.1a). 3. Reproduction by sporangiospores. The reproductive structure characteristic of Zygomycetes is the sporangiospore, an asexually produced spore which in genera of interest here is usually produced inside a sac, the sporangium, on the end of a long specialised hypha. Sporangiospores are produced very rapidly. From the food spoilage point of view, the outstanding properties of Zygomycetes are very rapid growth, especially in fresh foods of high water activity; inability to grow at low water activities (no Zygomycetes are xerophiles); and lack of resistance to heat and chemical treatments. From the food safety point of view, Zygomycetes have rarely been reported to produce mycotoxins.

3.4 Ascomycotina The subkingdom Ascomycotina is distinguished from Zygomycotina by a number of fundamental characters, the most conspicuous being the production of septate mycelium (Fig. 3.1b). Consequent on

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this, growth of fungi in this subkingdom is usually slower than that of Zygomycetes, although there are some exceptions. Fungi in the subkingdom Ascomycotina, loosely called ‘‘ascomycetes’’, characteristically produce their reproductive structures, ascospores, within a sac called the ascus (plural, asci, Fig. 3.2a, b). In most fungi, nuclei normally exist in the haploid state. At one point in the ascomycete life cycle, diploid nuclei are produced by nuclear fusion, which may or may not be preceded by fusion of two mycelia. These nuclei undergo meiosis within the ascus, followed by a single mitotic division and then differentiation into eight haploid ascospores. In most genera relevant to this work, asci can be recognised in stained wet mounts by their shape, which is spherical to ellipsoidal and smoothly rounded; size, which is generally 8–15 mm in diameter; and the presence when maturity approaches of eight ascospores tightly packed within their walls. At maturity asci often rupture to release the ascospores, which are thick walled, highly refractile, and often strikingly ornamented (Fig. 3.2c, d). Two other characteristics of asci are significant: generally they mature slowly, after incubation for 10 days or more at 258C, and they are usually borne within a larger, macroscopic body, the general term for which is ascocarp. Genera of interest here usually produce asci and ascospores within a spherical, smooth-walled body, the cleistothecium (Fig. 3.3a), or a body with hyphal walls, the gymnothecium (Fig. 3.3b). Ascospores are highly condensed, refractile spores, which are often resistant to heat, pressure

Fig. 3.1 (a) Nonseptate mycelium of Syncephalastrum racemosum; (b) septate mycelium of Fusarium equiseti

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Naming and Classifying Fungi

Fig. 3.2 Asci and ascocarps: (a) asci of Talaromyces species; (b) asci of Byssochlamys fulva; (c) ascospores of Eupenicillium alutaceum; (d) ascospores of Neosartorya quadricincta. Bars ¼ 5 mm

and chemicals. Almost all xerophilic fungi are ascomycetes. Besides their sexual spores, ascospores, ascomycetes commonly produce asexual spores. Formed after mitotic nuclear division, these spores are borne singly or in chains, in most genera of interest here from more or less specialised hyphal structures. The general term for this type of spore is conidium (plural, conidia), but other more specialised terms

exist for specific kinds of conidia. Along the evolutionary process, some Ascomycetes with welldeveloped asexual stages lost the ability to produce ascospores, and rely entirely on conidia for dispersal. Conidia, and the specialised hyphae from which they are borne, are astonishingly diverse in appearance. The size, shape and ornamentation of conidia and the complexity of the structures producing them

Fig. 3.3 (a) Cleistothecia of Eupenicillium; (b) gymnothecia of Talaromyces. SEM. Bars = 50 mm

3.7 Dual Nomenclature

provide the basis for classification of Ascomycetes that no longer produce the ascosporic (sexual) stage. Lacking ascospores, conidial fungi are not usually heat resistant, but conidia may be quite resistant to chemicals. Some conidial fungi are xerophilic.

3.5 Basidiomycotina The Subkingdom Basidiomycotina includes mushrooms, puffballs and the plant pathogenic rusts and smuts. Until recently it was not considered of any interest to the food mycologist. However, molecular studies indicate that the small brown species Wallemia sebi, long a curiosity because of its lack of resemblance to any other fungus, is a basidiomycete. It has no obvious phylogenetic affinity with any other genus and has now been classified in its own order, Wallemiales (Zalar et al., 2005). Only one other species of foodborne fungi, Trichosporonoides nigrescens Hocking and Pitt (1981), has a known affinity with this subkingdom.

3.6 The Ascomycete – Conidial Fungus Connection It was established more than a century ago that many fungal species carry the genetic information to produce both ascospores and conidia. These two kinds of spores are produced by different mechanisms and have different functions, so they are not always formed simultaneously. Not surprisingly, mycologists sometimes have given different generic and species names to a single fungus producing both an ascosporic and a conidial state. The usage of these names under the ICBN depends on the circumstances under which they were originally given. Some of these circumstances are discussed briefly below. The ascomycete state, now usually referred to as the teleomorph, is regarded by nomenclaturalists as the more important reproductive state, and the name applied to the teleomorph should be used when the ascomycete state is present. If the conidial state is also in evidence, the fungus is now a holomorph and is still correctly known by the teleomorph name. If the conidial state, known as the

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anamorph, has a separate name, this strictly speaking applies to the conidial state. It should be used only when the ascomycete state is absent, or to refer specifically to the conidial state if the ascomycete is present. However, the reader is warned that some anamorphic names are, and will continue to be, in common use for holomorphic fungi. Under the Articles of the ICBN, a generic name originally given to an anamorphic or conidial fungus cannot be used for a teleomorphic or ascomycetous fungus. For example, the name Penicillium, originally given to an anamorphic fungus with no known teleomorph, cannot be used for the teleomorphs later found to be produced by other Penicillium species. Such teleomorphs are classified in the genera Eupenicillium or Talaromyces, depending on whether ascospores are produced in cleistothecia or gymnothecia. Correct species names for the ascomycetous and conidial states of a single holomorphic fungus may or may not be the same, depending both on the circumstance in which the names were originally given, and on later synonymy. For example, Eupenicillium ochrosalmoneum Scott and Stolk and Penicillium ochrosalmoneum Udagawa refer to the teleomorph and anamorph of a single fungus. Udagawa (1959) described the anamorph; the teleomorph was later found, in the same isolate, by Scott and Stolk (1967). On the other hand, the anamorph of Eupenicillium cinnamopurpureum Scott and Stolk (1967) is Penicillium phoeniceum van Beyma (1933), with P. cinnamopurpureum Abe ex Udagawa (1959) as a synonym. Scott and Stolk (1967) found a teleomorph in Udagawa’s P. cinnamopurpureum; Pitt (1979b) later showed that this species was a synonym of the earlier P. phoeniceum. E. cinnamopurpureum, the first name applied to the teleomorph, is unaffected by this change in the anamorph name. In passing, note that ‘‘Abe ex Udagawa’’ indicates invalid (incomplete) publication of this species by Abe, with validation later by Udagawa. The species dates from the year of validation.

3.7 Dual Nomenclature An important point here is that some isolates of Penicillium phoeniceum regularly produce the teleomorphic state Eupenicillium cinnamopurpureum, while

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others, taxonomically indistinguishable, fail to produce a teleomorph at all. Because of this, it is essential to have a separate name for teleomorph and anamorph. The system of two names for a single fungus, known as dual nomenclature, has a place in the classification of fungi despite its apparent complexity. In the descriptions in later chapters, fungi for which both teleomorphs and anamorphs are known have both names listed. As noted above, if both states are found in a particular isolate, the teleomorph name is the more appropriate: to use that given to the anamorph is not incorrect, but this name is more sensibly applied to the conidial state only. Dual nomenclature would be relatively simple if the relationship between anamorph and teleomorph was always one to one. This is not the case. As has already been mentioned, species classified in Penicillium may produce teleomorphs in two genera, Eupenicillium and Talaromyces. On the other hand Talaromyces produces anamorphs in two genera, Penicillium and Paecilomyces. Aspergillus is the anamorph of eight or ten teleomorphic genera. Most teleomorph–anamorph relationships encountered in food mycology belong to the genera mentioned here. These relationships will be described where necessary under these particular genera.

3.8 Practical Classification of Fungi Fungi are classified in a vast array of orders, families, genera and species. Among natural organisms, the numbers of taxa of fungi are rivalled only by those of the flowering plants and insects. Estimates of fungal species range as high as 1.5 million; only 5% of this number have so far been described (Hawksworth, 1991). Many fungi are highly specialised. Some will grow only in particular environments such as soil or water; many are obligate parasites and require a specific host, such as a particular plant species, and will not grow in artificial culture; many grow only in association with plant roots. From the point of view of the food microbiologist, these kinds of fungi are irrelevant. In one sense, most fungi which spoil foods are also highly specialised, their speciality being the ability to obtain nutrients from, and hence grow on, dead, dormant or moribund plant material more or less regardless of source. The

3

Naming and Classifying Fungi

principal factors influencing food spoilage by fungi are physico-chemical and have already been outlined in Chapter 2. The point being made here is that food spoilage fungi are classified in just a few orders and a relative handful of genera. For this reason there is much to be said for food mycologists avoiding the use of a traditional, hierarchical classification as outlined above and employing a less formal approach to the identification of the fungi of interest to them. In the present work, this pragmatic approach has been followed as far as possible:

 The use of specialised terms has been kept to a 







minimum, while being cognisant of the need for clarity of expression. Hierarchical classification has been avoided as far as possible, consistent with retaining a logical approach to the presentation of fungi which are related or of similar appearance. Identification procedures used have been designed to be simple and comprehensible, avoiding the use of specialised equipment or procedures unavailable in the routine laboratory. To this end, identification of nearly all species included in this work is based entirely on inoculation of a single series of Petri dishes, incubation under carefully standardised conditions and examination by traditional light microscopy. A standard plating regimen has been used for the initial examination of all isolates (except yeasts), so that identification procedures can be carried out without foreknowledge of genus or even subkingdom. Cultural characters, which can be broadly defined as the application of microbiological techniques to mycology, have been used throughout.

The use of cultural characters has long been implicit in the study of fungi in pure culture on artificial substrates, especially in such genera as Aspergillus and Penicillium, genera of paramount importance in food spoilage. In Penicillium, cultural characters have been used as taxonomic criteria since the turn of the 20th century, but have assumed greater importance through the work of Pitt (1973, 1979b), who used the measurement of colony diameters, following incubation under standardised conditions, as a taxonomic criterion. The use of

3.8 Practical Classification of Fungi

pure culture techniques and growth data in fungal taxonomy is now widespread. Food microbiologists, the primary audience for this book, are familiar with cultural techniques and the use of a wide range of media and varied incubation conditions, so the authors make no apology for the taxonomic approach used in the present work. This approach is a logical extension of the system used by the first author in Penicillium taxonomy and which has been found to have a much broader applicability. In the field of mycology, different genera have been studied by many different people of varied backgrounds and for different reasons. Consequently, keys and descriptions have been based on a wide variety of media, often traditional formulations incorporating all sorts of natural products. This heterogeneity makes comparisons difficult

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and adds unnecessary complexity to the task of the nonspecialist confronted with a range of fungal genera. The approach used here has been to examine every isolate (excluding yeasts) by a single system: inoculation onto a standard set of Petri dishes and examination of them culturally and microscopically after 7 days incubation. Most of the genera and species included in this book can be identified immediately, at that point. Only in exceptional cases has it been found necessary to reinoculate isolates onto a further set of media in order to complete identification. The exceptional fungi are first the xerophiles, many of which grow poorly if at all on the standard media, and, second, genera such as Fusarium and Trichoderma, in which some species cannot readily be differentiated on the standard regimen. Details of the techniques used are given in Chapter 4.

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Chapter 4

Methods for Isolation, Enumeration and Identification

This chapter describes techniques and media suitable for the enumeration, isolation and identification of fungi from foods. Some techniques are similar to those used in food bacteriology; others have been developed to meet the particular needs of food mycology. Most of the media have been specifically formulated for foodborne fungi. The approach taken here is designed to provide a systematic basis for the study of food mycology. In 1984 a group of about 30 of the world’s foremost scientists in food mycology met in Boston, Massachusetts, USA, to hear and discuss a wide range of presentations that explored many aspects of methodology in food mycology. Agreement was reached on broad issues and areas requiring further work pinpointed. The proceedings were published as ‘‘Methods for the Mycological Examination of Food’’ (King et al., 1986). At a second workshop, held in Baarn, the Netherlands, in 1990, results of a number of collaborative studies on media and methods were presented and some standardised protocols developed. The proceedings, published as ‘‘Modern Methods in Food Mycology’’ (Samson et al., 1992), provided a comprehensive overview of current thinking in this field. The working group which organised those two workshops was then formalised as the International Commission on Food Mycology (ICFM), a commission under the auspices of the Mycology Division of the International Union of Microbiological Societies (IUMS). ICFM is dedicated to international standardisation of methods in food mycology. Subsequent ICFM workshops were held in Copenhagen, Denmark (1994), Uppsala, Sweden (1998), Samsø, Denmark (2003) and Key West,

Florida (2007). Papers from the third and fourth workshops were published in the International Journal of Food Microbiology and the proceedings of the fifth (Samsø) workshop were published as ‘‘Advances in Food Mycology’’ (Hocking et al., 2006a). The methodology described below is based on recommendations from ICFM and represents current thinking within the food mycology community. However, no formal endorsement from ICFM is implied.

4.1 Sampling It must be emphasised at the outset that results from mycological assays of foods are only as good as the samples used. However, sampling is beyond the scope of this text. Excellent treatises on sampling plans for food bacteriological purposes have been produced by the International Commission on Microbiological Specifications for Foods (ICMSF, 1986, 2002) and are generally applicable to food mycology.

4.2 Enumeration Techniques Quantification of the growth of filamentous fungi is more difficult than for bacteria or yeasts. Vegetative growth consists of hyphae, which are not readily detached from the substrate and which survive blending poorly. When sporulation occurs,

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_4,  Springer ScienceþBusiness Media, LLC 2009

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very high numbers of spores may be produced, causing sharp rises in viable counts, often without any great increase in biomass. The estimation of fungal growth or biomass is not easy, because no primary standard exists (such as cell numbers used for yeasts and bacteria). Although techniques for quantifying biomass have improved in recent years, most food laboratories continue to rely on viable counting (dilution plating) for detecting and quantifying fungal growth in foods. As well as dilution plating, a second standard method, known as direct plating, has been developed for estimating fungal numbers and growth in foods. Both methods are described in detail below. Techniques for biomass estimation will be discussed later.

4.2.1 Direct Plating Contributions at the international workshops mentioned above emphasised the use of direct plating as the preferred method for detecting, enumerating and isolating fungi from particulate foods such as grains and nuts. In direct plating, food particles are placed directly on solidified agar media. In most situations, particles should be surface disinfected before plating, as this removes the inevitable surface contamination arising from dust and other sources and permits recovery of the fungi actually growing in the particles. This process provides an effective measure of inherent mycological quality and permits assessment of the potential presence of mycotoxins as well. Surface disinfection should be omitted only where surface contaminants become part of the downstream mycoflora, for example, in grain intended for flour manufacture. Even here, surface disinfection before direct plating provides the most realistic appraisal of actual grain quality. Results from direct plating analyses are expressed as percentage infection of particles. The technique provides no direct indication of the extent of fungal invasion in individual particles. However, it is reasonable to assume that a high percentage infection is correlated with extensive invasion in the particles and a higher risk of mycotoxin occurrence.

4 Methods for Isolation, Enumeration and Identification

The standard protocol recommended by the ICFM (Hocking et al., 2006a, p. 344) is given below, with amplification where necessary. Surface disinfection. Surface disinfection is carried out by immersing particles in a chlorine solution. Household chlorine bleach, nominally 4–5% active chlorine, is effective. Dilute the chlorine 1 to 10 with water before use, to provide an approximately 0.4% solution. Immerse particles for 2 min, stirring occasionally, then drain the chlorine. Chlorine solutions are rapidly denatured by organic matter, so it is important to use a surplus of chlorine solution (10 times the volume of the particles) and to use the solution only once. This process is conveniently carried out in 250–500 ml beakers. Place 50 or more particles in the beaker and add chlorine. To dislodge air bubbles, immediately stir with a pair of forceps, leaving them in the solution, and preferably cover the beaker with a watch glass. The watch glass simplifies decanting of the chlorine, and the forceps, disinfected by the chlorine, may be used to plate the particles. Studies in our laboratory have shown that the treatment outlined here may be inadequate under some conditions. In commodities such as peanuts or maize where high levels of Aspergillus flavus or Penicillium species may be present, surface disinfection may be difficult. Here 2 min immersion in 70% ethanol followed by 2 min in 0.4% chlorine is recommended. Rinsing. After the chlorine is poured off, particles may be rinsed once with sterile water. Use a 1 min treatment, with stirring, then pour the water off. Again a watch glass should cover the beaker during this period, and the sterile forceps should be used for stirring. It is not clear whether rinsing fulfils any essential function. Early direct plating regimens used agents such as mercuric chloride for disinfection, and rinsing was essential to remove such toxic materials before they penetrated the particles too deeply. However, chlorine is effectively denatured by the particles and is believed to penetrate very little. In our opinion, the rinsing step can be omitted without loss of efficacy of the treatment, with savings in time and materials, and reduced risk of recontamination from the air. Plating. After disinfection and the optional rinse, particles should be plated onto solidified agar, at the rate of 6–20 particles per plate, depending on

4.2 Enumeration Techniques

particle size. Use the disinfected forceps. Plating should be carried out immediately: keep the watch glass on the beaker if this is not possible. Incubation. Incubate plates upright, under normal circumstances for 5 days at 258C. See more detailed notes below. Examination. After incubation, examine plates visually, count the numbers of infected particles and express results as a percentage. Differential counting of various genera is often possible. Correct choice of media, a stereomicroscope and experience will all assist in this process.

4.2.2 Dilution Plating Dilution plating is the appropriate method for mycological analysis of liquid or powdered foods. It is also suitable for grains intended for flour manufacture and other situations where total fungal contamination is relevant. Sample preparation. The two most common methods of sample preparation for dilution plating are stomaching and blending: stomaching is recommended by ICFM (King et al., 1986). The Colworth Stomacher (Sharpe and Jackson, 1972), or equivalent equipment (e.g. BagMixer1, Interscience, Saint Nom La Brete`che, France), is a very effective device for dispersing and separating fungi from finely divided materials such as flour and spices, and soft foods such as cheeses and meats, and its use is strongly recommended. Treatment time in the stomacher should be 2 min. Harder or particulate foods such as grains, nuts or dried foods, like dried vegetables, should be soaked before stomaching. Soaking times from 30 to 60 min are generally sufficient, but for extremely hard particles such as dried legumes, soaking for up to 3 h may be required. Comminution in a Waring Blender or similar machine is a suitable alternative for these types of samples and may give a more satisfactory homogenate. Blending times should not exceed 60 sec, as longer treatments may fragment mycelium into lengths too short to be viable or overheat the homogenate. The sample size used should be as large as possible. If a stomacher or BagMixer 400 is used, a sample size of 10–40 g is suitable.

21

Diluents. The recommended diluent is aqueous 0.1% peptone (Hocking et al., 2006a, p. 344), suitable for both filamentous fungi and yeasts. Saline solutions, phosphate buffer or distilled water are no longer recommended by ICFM as they may be deleterious to yeasts (Mian et al., 1997). The addition of a wetting agent such as polysorbate 80 (Tween 80) may be desirable for some products, but the natural wetting ability of the peptone is usually adequate. Special diluents may be necessary in some circumstances. If yeasts are to be enumerated from dried products or juice concentrates, the diluent should also contain 20–30% sucrose, glucose or glycerol, as the cells may be injured or be susceptible to osmotic shock. Dilution. Serial dilutions of fungi are carried out by the same procedures as those used in bacteriology, and the recommended dilution rate is 1:10 (=1+9). Fungal spores sediment more quickly than bacteria, so it is important to draw aliquots for dilution or plating as soon as possible, preferably within 1 min (Beuchat, 1992). Plating. Spread (surface) plating is recommended. When pour plates are used, fungi develop more slowly from beneath the agar surface and may be obscured by faster growing colonies from surface spores. Hence spread plating allows more uniform colony development, improves the accuracy of enumeration of the colonies and makes subsequent isolation of pure cultures easier. The optimum inoculum for surface plating is 0.1 ml. Best results will be obtained if plates are dried slightly before use. After adding the inoculum, spread it evenly over the agar surface with a sterile bent glass rod (‘‘hockey stick’’). Sterilise the rod by flaming it with ethanol before use. A plate spinner is a useful accessory. It is usually possible to enumerate plates with up to 150 colonies, but if a high proportion of rapidly growing fungi are present, the maximum number which can be distinguished with any accuracy will be lower. Because of this restriction on maximum numbers, it may be necessary on occasion to accept counts from plates with as few as 10–15 colonies. Clearly, such limitations on numbers per plate and the overgrowth of slow colonies will result in higher counting errors than are usually achieved with bacteria or yeasts. Enumerating yeasts is less difficult. In the absence of filamentous fungi, from 30 to 300 colonies per

22

plate can be counted and errors will be comparable with those to be expected in bacterial enumeration. Incubation. The standard incubation conditions are 258C for 5 days. However, other conditions may be more suitable in some circumstances (see notes below). Reporting results. As in bacteriology, results from dilution plating are expressed as viable counts per gram of sample. Note that such results are not directly comparable with those obtained from direct plating and may not offer a direct indication of the extent of fungal growth.

4.2.3 Incubation Conditions As noted above, the standard incubation conditions specified by ICFM are 258C for 5 days (Hocking et al., 2006a). Undoubtedly, 258C is the most suitable temperature for routine work in temperate to subtropical environments. Few if any common fungi are sensitive to this temperature, even those which grow readily under refrigeration. Higher temperatures are unacceptable in the temperate zone: 308C is close to the upper limit for growth of some important Penicillium species. In tropical regions, incubation at 308C is recommended as a more realistic temperature for enumerating fungi from commodities stored at ambient temperatures. In cool temperate regions such as Europe, 228C may be a more suitable incubation temperature. When used for fungi, Petri dishes should be stored upright. The principal reason is that some common fungi can shed large numbers of spores during handling, which in an inverted dish will be transferred to the lid. Reinversion of the Petri dish for inspection or removal of the lid may liberate spores into the air or onto benches and cause serious contamination problems.

4.3 Sampling Surfaces Methods are outlined below for directly sampling the mycoflora of surfaces of commodities such as fruits, meats, cheeses, salamis and dried fish and also packaging materials, machinery and walls.

4 Methods for Isolation, Enumeration and Identification

The techniques are based on those described by Langvad (1980) for studying the fungal flora of leaves. If samples are particulate, or can be cut up, sterile forceps can be used to press pieces of a suitable size (up to about 10 mm2) onto a suitable medium in a Petri dish. The sample is then removed, leaving an impression, and any spores or mycelium transferred will form colonies within a few days. This technique is known as press plating. For packaging materials such as cardboard, an alternative method is to cut a piece which will fit in a standard Petri dish. The dish is prepared by adding a sterile filter paper moistened with 10% glycerol and then placing a bent glass rod on it as a separator. After adding the sample, a thin layer of an appropriate agar medium is poured over its surface. To reduce evaporation, the dish should be sealed with Parafilm or a similar material or placed in a polyethylene bag before incubation at 258C for a few days. If contamination levels are not too heavy, the number and types of moulds present can be effectively estimated by this method. Colonies may be subcultured for identification. For sampling walls or other surfaces, or for nondestructive sampling, impressions may be taken using adhesive tape. Carefully handled, tape coming from the roll will be virtually sterile. Press a short length of tape firmly onto the surface to be sampled, adhesive side down, then transfer it, still with the same side down, onto a suitable growth medium. After 1–2 days incubation at 258C, the tape may be removed to allow development and sporulation of colonies. Surface sampling techniques for assessment of sanitation in food production and processing areas are discussed by Evancho et al. (2001). Surfaces may be sampled by using sterile swabs or by agar contact methods. The swab method involves rubbing a moistened sterile cotton swab over the test surface and placing the swab in a dilution bottle to be subsequently diluted and plated on appropriate media. Agar contact plates, also known as RODAC (replicate organism direct agar contact) plates, are restricted to use on smooth or semi-smooth flat surfaces. An alternative to agar contact plates is the agar slice technique, where agar is filled into a syringe-like apparatus or into an artificial sausage casing. The solidified agar can be pushed out onto

4.5 Isolation Techniques

the surface to be sampled, then the portion making contact sliced off with a sterile scalpel or wire and placed in a Petri dish for incubation.

4.4 Air Sampling Air sampling in the food processing environment is discussed by Evancho et al. (2001). The simplest method of air sampling is by sedimentation or settle plates. A Petri dish containing an appropriate agar medium is exposed to the atmosphere for a fixed period of time (usually 15–60 min), then closed and incubated at 258C (Samson et al., 2004a). This method can be useful in food production areas, as it gives a direct indication of the number and types of fungi likely to come into contact with exposed product. However, the settle plate technique lacks precision, and volumetric air sampling is a much more reliable indicator of air quality. A number of air sampling devices are commercially available. Of these, the Anderson sampler (either the two-stage or six-stage model; Anderson Instruments Inc, Atlanta, GA, USA) is probably the best, giving accurate and consistent results (Buttner and Stetzenbach, 1993). However, the Anderson sampler is expensive and requires mains power or a large battery for operation. In factory situations, the dry cell battery-operated Biotest

23

Hycon RCS and RCS High Flow centrifugal air samplers (Biotest, Solihull, UK) and the MAS-100 air sampler (Merck, Darmstadt, Germany) are more convenient, as they are small and readily portable. The Biotest RCS Plus sampler was reported to give comparable results to larger and more sophisticated machines, but its sampling efficiency gradually decreased for particle sizes below 4 mm (Benbough et al., 1993).

4.5 Isolation Techniques The term ‘‘isolation’’ is used here in its strict sense: the preparation of a pure culture, free from any contamination and ready for identification.

4.5.1 Yeasts Streaking techniques commonly used for bacterial purification are equally suitable for the isolation of yeasts (Fig. 4.1). A method widely used by yeast specialists is to disperse a portion of a colony in 2– 3 ml of sterile water, then streak a single loop of this suspension over the whole surface of a plate, moving the loop slowly down from top to bottom while simultaneously moving it rapidly across the plate

Fig. 4.1 Petri dishes of yeasts growing on malt extract agar, showing a streaking method suitable for producing isolated colonies

24

from side to side. After suitable incubation, wellseparated single colonies should appear in the lower half of the plate (Fig. 4.1). If all of these single colonies appear to be of similar size and appearance (taking into account the effect of crowding), the culture may be judged to be pure. Microscopic checks of some single colonies are also desirable. Disperse a needle point of cells from a colony in a drop of water, add a cover slip and examine by bright field illumination at about 400. Cell outlines will be clearly visible. Note that, unlike those of bacteria, yeast cell sizes often vary considerably in a pure preparation. Purity is indicated not so much by uniformity of cell size within a preparation as by similarity in microscopic cell appearance from colony to colony. When a culture is considered to be pure, streak it onto an appropriate slant.

4.5.2 Moulds Streaking techniques are ineffective for filamentous fungi and are not recommended. Isolation depends on picking a small sample of hyphae or spores – judged to be pure by eye, by hand lens or preferably under the stereomicroscope – and placing this sample on a fresh plate as a point inoculum. Purity is subsequently judged by uniformity in appearance of the colony which forms after incubation. The appearance of a mixed culture depends on the growth rates of the fungi present. If rates are diverse, a mixed culture is often indicated by a clump of dense hyphae at the inoculum point, surrounded by looser wefts of spreading hyphae. With fungi of approximately equal growth rates, mixtures are often indicated by colonies with sectoring growth: sectors will show differences in mycelial, spore or reverse colours or in radial growth rates. The simplest starting place for isolating fungi is an enumeration plate with well-separated colonies. Use a needle of platinum or nichrome, preferably cut to a chisel point with a pair of pliers, or a steel sewing needle. Sterilise it by heating, then plunge the tip into cold agar and leave until cool – with nichrome or steel this will require several seconds. With the tip of the cold, wet needle pick off a few spores or a tuft of mycelium – just enough to be

4 Methods for Isolation, Enumeration and Identification

visible – and inoculate a single point on a plate or slant. The same procedure can be applied to mixed cultures arising from direct plating or surface sampling techniques. It is advisable to keep notes on the appearance of the colony area sampled, as this will give an indication of whether the culture which grows up is the same as that intended to be isolated. It is generally easy to isolate rapidly growing fungi from those which grow more slowly. The outermost hyphal tips are usually free of contamination. The reverse process is often much more difficult. The isolation of slowly growing fungi in the presence of rapidly growing ‘‘weeds’’ often requires skill, patience and ingenuity. It is desirable to watch the point inocula daily over several days at least, because a particular stage in the life cycle may give some advantage. The slow colony may germinate more rapidly, or have a sector accessible to a needle, or spore more freely. The use of higher or lower incubation temperatures, or media of low aw or low nutrient status, or the addition of dichloran or rose bengal (see below) may all be of value in this process. Freeing fungi from bacteria, long considered to be a very difficult procedure, has been greatly simplified in recent years with the advent of media containing antibiotics. With use of the media recommended in the next section, bacterial contamination of cultures should be a rare event. When a pure colony is obtained, it should be inoculated onto a slant of an appropriate medium and incubated until ready for identification. Again, moulds should always be inoculated at a single point, preferably near the centre of the slant. This permits the best colony development and sporulation in most fungi.

4.5.3 Short Term Storage For short to medium-term storage, fungi are usually stored on slants, most commonly in McCartney or Universal bottles which have the advantages of being free standing and having caps that can be sealed to retard drying during storage. During incubation and until colonies are fully mature, however, caps must be kept loose on slants in bottles. Moulds

4.6 Choosing a Suitable Medium

25

require free access to oxygen for typical growth and sporulation. Oxygen starvation during growth will at best lead to retarded sporulation or at worst death of the culture. Long-term preservation of fungi is dealt with later in this chapter.

4.6 Choosing a Suitable Medium Food laboratories often rely on a single all-purpose medium to produce a ‘‘yeast and mould’’ count in everything from raw material to finished product. But just as the food bacteriologist uses selective media for particular purposes, so too food mycologists are developing a range of media suited to specific applications. It is plainly unrealistic to expect a single medium to answer all questions about mould and yeast contamination in all foods. The fungi that spoil meats or fresh vegetables are not the same as those that grow on dried fish. Although often used for this purpose, very dilute media such as potato dextrose agar are of little or no value for enumerating fungi from dried foods.

The most important division in types of enumeration media lies between those suitable for high water activity foods, such as eggs, meat, vegetables and dairy products, and those suited to the enumeration of fungi in dried foods. The most suitable media for dried foods depend on the type of food, the major categories being foods low in soluble solids such as cereals, high sugar foods such as confectionery and dried fruit, and salt foods. A second consideration lies in whether the primary interest is in moulds, or yeasts, or both; and a third concerns the presence or absence of preservatives. Finally, media are available for specific mycotoxigenic fungi, notably Aspergillus flavus and related species and Penicillium verrucosum plus P. viridicatum. An overview of media considered most suitable for particular purposes is given in Table 4.1. The table is derived from Pitt and Hocking (1997), together with recent recommendations from ICFM (Samson et al., 1992; Hocking et al., 2006a).

Table 4.1 Recommended media for fungal detection, enumeration and isolationa Type of food

Selecting for

Medium

Remarks

Fresh foods: milk and milk products, fruit, cheese, sea foods

Moulds Yeasts General

DRBC TGY, MEA, OGY DRBC

Blend (where necessary) and dilution plate

Freshly harvested grains, nuts

General Dematiaceous Hyphomycetes Fusarium Yeasts

DRBC DRBC, CZID CZID TGY, MEA, OGY

Direct plate Direct plate Direct plate Dilution plate

Fruit juices, fresh

Yeasts

TGY, MEA, OGY

Dilution plate

Fruit juices, preserved

Preservative resistant yeasts

TGYA, malt acetic agar

Dilution plate

Fruit juices, to be pasteurised, or pasteurised products

Heat resistant moulds

PDA, MEA

Special protocol

Fruit juice concentrates

Xerophilic yeasts

MY50G

Special diluents

Dried foods in general

General

DG18

Direct plate

Stored cereals, nuts

General Dematiaceous Hyphomycetes Fusarium

DG18 DRBC, CZID CZID

Direct plate Direct plate Direct plate

26

4 Methods for Isolation, Enumeration and Identification

Table 4.1 Recommended media for fungal detection, enumeration and isolationa (continued) Type of food

Selecting for

Medium

Remarks

Grain for milling into flour

General

DG18

Stomach or blend and dilution plate

Dried fruit, confectionery, chocolate, etc.

Xerophilic moulds and yeasts

MY50G

Direct plate

Fastidious xerophiles – in presence of Eurotium spp.

MY50G MY70GF

Direct plate Direct plate

General

DG18

Halophilic xerophiles

MY5-12, MY10-12

Direct plate or press plate Direct plate or press plate

Fungi producing aflatoxins

AFPA

Direct or dilution plate

DRYS

Direct or dilution plate

Salt foods, e.g. salt fish

General

General Fungi producing ochratoxins a For medium acronyms, see Section 4.6.

4.6.1 General Purpose Enumeration Media To be effective, a general purpose enumeration medium must fulfil several requirements (Pitt, 1986). As these are sometimes overlooked, they are listed here:

 to inhibit bacterial growth completely, without  

  

affecting growth of foodborne fungi (filamentous or yeasts); to be nutritionally adequate and support the growth of fastidious fungi; to suppress the growth of rapidly spreading fungi, especially the Mucorales, but not to prevent their growth entirely, so they too can be enumerated; to slow radial growth of all fungi, to permit counting of a reasonable number of colonies per plate, without inhibiting spore germination; to promote growth of relevant fungi; and to suppress growth of soil fungi or others generally irrelevant in food spoilage.

Fulfilling the above requirements necessitates the use of potent inhibitory compounds, and there is sometimes a fine line between inhibition of undesirable microorganisms and suppression of growth of those being sought. Modern fungal enumeration media rely on the use of antibiotics at neutral pH

for the inhibition of bacteria. Such media allow better recovery of moribund and sensitive fungi than the acidified media commonly used in the past. For many years rose bengal has been added to media to slow colony spread, while more recently 2,6-dichloro-4-nitroaniline (dichloran) has been added to inhibit rapidly spreading moulds. Many common spoilage fungi, Aspergillus and Penicillium species in particular, develop better on media with adequate nutrients. Low nutrient media of very high aw, such as potato dextrose agar, have lost favour because they are selective against some species in these genera. The media described below are considered to be the most satisfactory general purpose enumeration media available at this time (Hocking et al., 2006a). Formulations are given in the Media Appendix. Dichloran rose bengal chloramphenicol (DRBC) agar. DRBC (King et al., 1979, Pitt and Hocking, 1997) is recommended for both moulds and yeasts. It is particularly suited to fresh and high aw foods (Hocking et al., 1992). This medium contains both rose bengal (25 mg/kg) and dichloran (2 mg/kg), which restrict colony spreading without affecting spore germination unduly. Compact colonies allow crowded plates to be counted more accurately. This combination of inhibitors also effectively restricts the rampant growth of most of the common mucoraceous fungi such as Rhizopus and

4.6 Choosing a Suitable Medium

27

Fig. 4.2 Petri dishes of (a) DRBC and (b) RBC showing effective control of Rhizopus growth by rose bengal and dichloran in DRBC

Mucor (Fig. 4.2), although it does not completely control some other troublesome genera such as Trichoderma. In routine use, it is recommended that DRBC plates be incubated away from light at 258C for 5 days. Dichloran 18% glycerol (DG18) agar. Hocking and Pitt (1980) developed DG18 to be selective for xerophilic fungi from low moisture foods such as stored grains, nuts, flour and spices. DG18 was designed for enumeration of a range of nonfastidious xerophilic fungi and yeasts. However, it has been shown since that it supports growth of the common Aspergillus, Penicillium and Fusarium species, as well as most yeasts, and many other common foodborne fungi. DG18 can now be described as a general purpose medium with emphasis on enumeration of fungi from dried foods. DG18 is also recognised to be a very useful medium for enumeration of airborne fungi (Wu et al., 2000; Samson et al., 2004a). It is an effective inhibitor of Mucoraceous fungi, and bacteria are totally suppressed. However, growth of Eurotium species (previously known as ‘‘the Aspergillus glaucus group’’) is still somewhat too rapid, and colonies may have diffuse margins. The addition of detergent to DG18 has been reported to be an improvement in this respect (Beuchat and de Daza, 1992). Although DG18 is a satisfactory isolation medium for Eurotium species, it is not suitable for their

identification. Eurotium species are usually identified on Czapek yeast extract agar with 20% sucrose (CY20S), described later in this chapter. Other general purpose media. Under circumstances where rapidly spreading moulds do not cause problems, two alternative general purpose enumeration media are satisfactory. These are rose bengal chloramphenicol agar (RBC; Jarvis, 1973), from which DRBC was developed, and oxytetracycline glucose yeast extract agar (OGY; Mossel et al., 1970). OGY has been found to be very suitable for yeasts in the absence of moulds (Andrews, 1992a). Most of the media discussed above are available in ready to use dehydrated form from media suppliers such as Oxoid, Difco, BBL, etc.

4.6.2 Selective Isolation Media Although considerable progress has been made in the past 20 years, the formulation of selective media for foodborne fungi still requires a great deal of research. The availability of effective media can greatly simplify the isolation and identification of significant food spoilage and mycotoxigenic fungi. Most attention has been paid so far to the requirements of xerophilic fungi because of their failure to develop on standard high aw media. For mycotoxigenic fungi, satisfactory media exist only for

28

Aspergillus flavus and closely related species, Penicillium verrucosum and P. viridicatum, and the genus Fusarium. These media are considered below. Media for Aspergillus flavus and related species. Bothast and Fennell (1974) developed Aspergillus differential medium after finding that Aspergillus flavus and A. parasiticus produced conspicuous, diagnostic orange yellow colours in the colony reverses in the presence of an appropriate nitrogen source and ferric salts. Few other fungi produced a similar colouration. It was later shown that these species produce aspergillic acid or noraspergillic acid, which react with ferric ammonium citrate to form the colour complex (Assante et al., 1981). However, development of the orange yellow reverse pigment is not indicative of aflatoxin production. This medium was reformulated by Pitt et al. (1983) to produce Aspergillus flavus and parasiticus agar (AFPA). As well as more effective concentrations of the active ingredients, AFPA contains dichloran and chloramphenicol to inhibit spreading fungi and bacteria, respectively. When incubated at 308C for 42–48 h, colonies of Aspergillus flavus, A. parasiticus and A. nomius are distinguished by bright orange yellow reverse colours. Only A. niger can be a source of error: it grows as rapidly as A. flavus and sometimes produces a yellow, but not orange, reverse. After 48 h A. niger colonies begin production of their diagnostic black or dark brown heads, which provide a ready distinction from A. flavus. Prolonged incubation of AFPA, beyond 4 days, is not recommended because A. ochraceus and closely related species may also produce a yellow reverse after this time. AFPA is recommended for the detection and enumeration of potentially aflatoxigenic fungi in nuts, maize, spices and other commodities. Its advantages include rapidity, as 48 h incubation is usually sufficient, specificity, and simplicity, as little skill is required in interpreting results. In consequence, it can be a simple, routine guide to possible aflatoxin contamination (Pitt, 1984). This medium is also very effective for enumerating A. flavus in soils, where levels down to 5 spores per gram can be detected. Where soils carry a heavy bacterial load, doubling the quantity of chloramphenicol or adding other antibiotics may be of value. Detection of aflatoxin production using Coconut Cream Agar. Dyer and McCammon (1994) developed

4 Methods for Isolation, Enumeration and Identification

Coconut Cream agar (CCA) to detect aflatoxin production in Aspergillus flavus and A. parasiticus. CCA can be made using any brand of commercial canned coconut cream (available from Asian food stores in many places). Dilute 50:50 with water, add agar (1.5%) and autoclave. Inoculate solidified plates with up to four colonies (picked with a wet needle) and incubate at 308C for 5–7 days. Examine, reverse upmost, under long wave length UV light. Colonies producing aflatoxins will fluoresce bluish white or white, especially in the centres. Ignore fluorescence from the Petri dish itself. Use an uninoculated coconut cream agar plate as a control (Dyer and McCammon, 1994). Plates inoculated with known nontoxigenic and toxigenic strains are also useful controls. A. parasiticus isolates almost always produce aflatoxins. Media for fungi producing ochratoxin A. Although ochratoxin A was first described from Aspergillus ochraceus, recent molecular studies indicate that A. westerdijkiae is the major ochratoxin A producing species in Aspergillus Section Circumdati (Frisvad et al., 2004). Aspergillus carbonarius is also an important source of ochratoxin A, particularly in grape products (Abarca et al., 2004; Leong et al., 2006a and references therein). However, there are no selective or indicative media for these fungi. When DG18 is used as the isolation medium, selection of colonies with light brown (ochre) or black sporulation is a good starting point for detecting A. westerdjikiae or A. carbonarius respectively. Coconut cream agar (Dyer and McCammon, 1994) can also be used to screen for ochratoxin production in A. carbonarius and species in Aspergillus Section Circumdati (Heenan et al., 1998). Plates are best incubated at 258C rather than 308C for potentially ochratoxigenic species of Aspergillus. Penicillium verrucosum is the most important ochratoxin a producer in the genus Penicillium (Pitt, 1987), although P. nordicum can also produce this mycotoxin (Frisvad and Samson, 2004). Frisvad (1983) developed Dichloran rose bengal yeast extract sucrose agar (DRYS) for selective enumeration of P. verrucosum. P. nordicum, P. viridicatum and P. aurantiogriseum are also selected by DRYS. P. verrucosum and P. nordicum produce ochratoxin A, P. verrucosum also produces citrinin, while the latter two species produce xanthomegnin and viomellein. According to Frisvad (1983), P. verrucosum colonies on DRYS have a violet brown reverse, and

4.6 Choosing a Suitable Medium

the latter two species produce yellow colonies with a yellow reverse. The incubation regimen recommended by Frisvad (1983) is 7–8 days at 208C. Subsequently, Frisvad et al. (1992) developed dichloran yeast extract sucrose 18% glycerol agar (DYSG) on which P. verrucosum produces a red brown reverse. Because of its lower aw, DYSG inhibits rapidly growing fungi such as Rhizopus and Mucor more effectively than DRYES. In a study of 76 samples of wheat, rye and barley, Lund and Frisvad (2003) found that DYSG was more effective than DRYES for screening grain samples for the presence of P. verrucosum and potential ochratoxin A contamination. Media for Fusarium species. Dichloran chloramphenicol peptone agar (DCPA; Andrews and Pitt, 1986) can be used to isolate Fusarium species from grains and other substrates. The medium was developed from Nash-Snyder medium (Nash and Snyder, 1962) a medium for enumeration of Fusarium species from soils. DCPA uses a low level of dichloran as a substitute for the high level of pentachloronitrobenzene (PCNB), and chloramphenicol rather than the antibiotic mixture used by Nash and Snyder (1962). When Fusarium species are dominant, DCPA is effective for their isolation from grains, animal feeds and soil. However, DCPA has been found to be less effective in mixed populations. DCPA is a very useful medium for the identification of Fusarium species because it often induces the abundant formation of macroconidia (Hocking and Andrews, 1987). A much more stringent medium, effective for isolation of most Fusarium species occurring in foods, is Czapek iprodione dichloran agar (CZID; Abildgren et al., 1987). As well as dichloran, this medium contains the fungicide iprodione. CZID is highly selective for Fusarium species, and is probably the best medium for Fusarium enumeration and isolation. CZID is suitable for isolation of Fusarium species by direct plating of surface disinfected grains and other commodities, or dilution plating of homogeneous samples such as flour. Some questions remain concerning whether CZID may be too selective and not support growth of all foodborne Fusarium species. However, the common species all grow well. Castella´ et al. (1997) developed a Fusarium selective medium (MGA 2.5) using 2.5 mg/L malachite green as the selective agent. They reported that MGA 2.5 was more selective than Nash-Snyder medium as it did not allow development of colonies of other fungal genera.

29

Bragulat et al. (2004) compared the efficacy of MGA 2.5 medium with DCPA, CZID and several other Fusarium media, using pure cultures of twelve Fusarium species commonly found in foods as well as naturally contaminated samples. They reported that there was no statistically significant difference in colony counts of the Fusarium spp. tested, but that colonies on MGA 2.5 were smaller than the other media. MGA 2.5 did not allow growth of other fungi such as Zygomycetes and yeasts from naturally contaminated samples, thus providing better selectivity than the other media. Media for dematiaceous Hyphomycetes. Although designed primarily for Fusarium isolation, CZID has been found in our laboratory to be very useful for isolating dematiaceous Hyphomycetes, provided that iprodione is added at half the usual concentration, as full strength iprodione tends to restrict colony diameters too severely. Alternaria, Bipolaris, Curvularia, Stemphylium and Ulocladium species will grow and sporulate well when incubated for 5 days at 258C with a 12 h photoperiod. Drechslera species will grow but will not sporulate on this medium. DRBC is also of value for isolating these fungi, but some species do not sporulate readily on it. DCPA, originally developed for isolation of Fusarium, may also be used as an isolation medium for the dematiaceous Hyphomycete genera mentioned above. Alternaria, Curvularia and related genera usually grow rapidly and sporulate well on DCPA. However, fungi should not be maintained or stored on DCPA for more than two weeks, as ammonia is produced by aging cultures. Media for xerophilic fungi. Xerophilic fungi are of great importance in food spoilage, and hence media and techniques for their enumeration and isolation have received much attention. Xerophiles range from those which grow readily on normal media and which are only marginally xerophilic, such as many Aspergillus and Penicillium species, to those, such as Xeromyces bisporus, which will not grow at all on normal (high aw) media. It is not surprising that no single medium is suitable for quantitative estimation of all xerophilic fungi found causing food spoilage. As noted earlier, DG18 was developed as a general medium for xerophiles, and remains the medium of choice for this purpose. DG18 should be used in any general examination of the mycoflora of dried foods. Media for extreme xerophiles. Fungi discussed here include Xeromyces bisporus, xerophilic

30

Chrysosporium species, Eremascus species, Eurotium halophilicum and the halophilic xerophiles Polypaecilum pisce and Basipetospora halophila. These extreme xerophiles grow slowly if at all on DG18 and are quickly overgrown by rapidly spreading xerophiles such as common Eurotium species. Such species require special media and techniques. Since the early 1950s, studies on extremely xerophilic fungi have been carried out in our laboratory (Scott, 1957). A very wide variety of media have been used for their cultivation. The most effective of these, suitable for all except the halophilic xerophiles, is malt extract yeast extract 50% glucose agar (MY50G; Pitt and Hocking, 1997). Isolation techniques for extreme xerophiles. As well as special media, extreme xerophiles require special techniques for isolation. If a low aw commodity such as ground spice, dried fruit, fruit cake, confectionery or dried fish shows signs of white mould growth, it is likely that the fungus responsible is an extreme xerophile. Fungi of this kind are usually sensitive to diluents of high aw and often cannot be isolated by dilution plating. For direct plating, a convenient technique is to place small pieces of sample, without surface disinfection, onto a rich, low aw medium, such as MY50G. Alternatively, examine the food under the stereomicroscope, which will often provide useful information in any case, and use an inoculating needle to pick off pieces of mycelium or spores from the surface of the spoiled food. Place these pieces (three to six per plate) directly onto MY50G. Colonies should develop after 1–3 weeks incubation at 258C. Quite frequently extreme xerophiles will be present in pure culture and can be readily isolated. Examination with the stereomicroscope will usually indicate whether this is the case. Selection of growth which appears to cover the range of types seen will assist isolation of the principal fungi present. Direct sampling by the press plating technique described earlier in this chapter can be useful for isolating xerophiles. Sometimes extreme xerophiles are accompanied by Eurotium species, also capable of growth at very low aw. The latter are identifiable under the stereomicroscope by their Aspergillus heads. Under these circumstances, isolation of extreme xerophiles is more difficult. A medium of sufficiently low aw to slow the growth of Eurotium species is necessary.

4 Methods for Isolation, Enumeration and Identification

The most satisfactory medium is malt extract yeast extract 70% glucose fructose agar (MY70GF). MY70GF is of similar composition to MY50G, except that it contains equal parts of glucose and fructose to prevent crystallisation of the medium at the concentration used (70% w/w). It is made in a similar manner to MY50G. Growth of even the extreme xerophiles on MY70GF is extremely slow, and plates should be incubated for at least 4 weeks at 258C. Once growth is apparent, pick off small portions of colonies and transfer them to MY50G, to allow more rapid growth and sporulation. Media for halophilic xerophiles. Some xerophilic fungi from salted foods such as salt fish grow more rapidly on media containing NaCl and hence are correctly termed halophilic xerophiles. Malt extract yeast extract 5% salt 12% glucose agar (MY5-12) and malt extract yeast extract 10% salt 12% glucose agar (MY10-12) are suitable for these fungi. Techniques for enumeration and isolation are similar to those described above for other extreme xerophiles.

4.6.3 Techniques for Yeasts The simplest enumeration and growth medium for most food spoilage yeasts is malt extract agar (MEA). Although originally introduced as a growth medium for moulds, its rich nutritional status makes it very suitable for yeasts, and its relatively low pH (usually near 5.0) reduces the possibility of bacterial contamination. More recently, tryptone glucose yeast extract agar (TGY) has been recommended for enumeration of yeasts (Hocking et al., 1992, 2006a). Due to a higher glucose concentration (10%) and higher pH (5.5–6.0), this medium is more effective in recovery of stressed yeast cells, and colony development is usually faster than on MEA. However, its higher pH means that an antibiotic should be incorporated for enumeration of yeasts from food samples which may contain bacteria. Recommended antibiotics are chloramphenicol or oxytetracycline at the concentrations used in DRBC and OGY. Both MEA and TGY are suitable for enumeration of yeasts in products such as fruit juices, fruit purees and yoghurt, where moulds are usually present only in low numbers.

4.6 Choosing a Suitable Medium

If large numbers of moulds are present, often the case with solid products such as cheese, a general purpose enumeration medium such as DRBC or DG18 should be used. For circumstances where yeast enumeration remains difficult on media such as DRBC, a valuable technique is that of de Jong and Put (1980), which involves a 3-day anaerobic incubation. They recommended using pour plates or streak plates of mycophil agar (BBL Microbiological Systems, Cockeysville, Maryland, USA), but malt extract agar should be equally suitable. Antibiotics may be added if necessary. Plates are incubated at 258C for 3 days in an anaerobic jar, then taken out and incubated aerobically for another 2 days at 258C. As many spoilage yeasts are able to grow anaerobically, colonies will commence development without the possibility of overgrowth by moulds. This technique is unlikely to be suited to the enumeration of yeasts on surfaces of fresh fruit or vegetables, as yeasts which colonise these surfaces are often strict aerobes. Enrichment for yeasts in liquid products. In liquid food products, low numbers of yeasts may be difficult to detect, but may have a serious potential to cause spoilage. Enrichment techniques are the only satisfactory way of monitoring products in these circumstances. For products or raw materials free from suspended solids and of low viscosity, standard membrane filtration techniques are a satisfactory method for detecting low numbers of yeasts. The filter can be placed directly onto a suitable medium such as MEA or TGY and staining can be carried out subsequently. Centrifugation may also be used, but has the disadvantage that only relatively small volumes of product can be screened. If products or raw materials are viscous, of low aw, or contain pulps and cannot be filtered efficiently, other enrichment techniques are needed. In many cases, the best enrichment medium is the product itself, diluted 1:1 with sterile water. A 1:1 dilution increases the aw of juice concentrates or honey to a level which will allow growth of potential spoilage yeasts without causing a lethal osmotic shock to the cells. If the product contains preservative, dilution will lower the concentration and permit cells to grow. To detect low numbers of spoilage yeasts in cordials, fruit juice concentrates and similar materials, simply decant half the product from the container

31

and replace it with sterile water. Leave the cap loose, incubate at room temperature or 308C and watch for evidence of fermentation. Shaking the container daily will help to detect gases resulting from fermentation. Classical enrichment techniques used in bacteriology can also be used for yeasts. TGY broth has been used very successfully in our laboratory for enrichment of low numbers of yeasts in liquid products. Add 10 ml of product to 90 ml TGY broth and incubate at 258C for 3–4 days, or 308C for 2–3 days. Look for signs of fermentation and streak out onto TGY agar. Detection of preservative resistant yeasts. A few species of yeasts are able to grow in products containing preservatives such as sorbic, benzoic and acetic acids or sulphur dioxide. The most important of these is Zygosaccharomyces bailii. The simplest and most effective way to screen for preservative resistant yeasts is to spread or streak product onto plates of malt acetic agar (MAA), which is MEA with 0.5% acetic acid added (Pitt and Richardson, 1973) or TGY with 0.5% acetic acid added (TGYA, Hocking, 1996). MAA and TGYA are made by adding glacial (16 N) acetic acid to melted and tempered basal medium to give a final concentration of 0.5%. Mix and pour immediately. These media cannot be held molten for long periods or remelted because of their low pH (approximately 3.2 and 3.8 for MAA and TGYA, respectively). The acetic acid does not need sterilisation before use. MAA and TGYA are suitable media for monitoring raw materials, process lines and products containing preservatives for resistant yeasts. They are also effective for testing previously isolated yeasts for preservative resistance. Erickson (1993) developed a selective medium for Zygosaccharomyces bailii. Zygosaccharomyces bailii medium (ZBM) is based on Sabouraud dextrose agar amended with fructose, NaCl, tryptone, yeast extract and trypan blue dye, then acetic acid (0.5%) and potassium sorbate (0.01%) are added to make the medium selective. It is designed as a plating medium for detection of Z. bailii in acidified ingredients in conjunction with hydrophobic grid membrane filtration (Erickson, 1993). When compared with MAA and TGYA in an interlaboratory study, ZBM was found to be highly selective for Zygosaccharomyces bailii, to the exclusion of other important preservative resistant yeasts such as Schizosaccharomyces pombe and Pichia

32

membranaefaciens. In addition, recovery of Z. bailii cells sublethally injured by lyophilisation was significantly lower on ZBM than on TGYA or MAA (Hocking, 1996). Its high selectivity and complex formulation make ZBM unsuitable for routine laboratory use as a medium for detection of preservative resistant yeasts. Enrichment of preservative resistant yeasts. A technique capable of detecting yeast numbers as low as 1 cfu/ml within 4 days has been developed in our laboratory (Hocking et al., 1996). This method is particularly suitable for the detection of Zygosaccharomyces bailii in raw materials or finished product, but can also be used for the detection of Schizosaccharomyces pombe, Pichia membranaefaciens and other species of preservative resistant yeasts. The method involves a 2to 3-day enrichment step followed by a plating step with a further 2 days of incubation. Triplicate 20 ml tryptone glucose yeast extract (TGY) broths containing 0.5% acetic acid (TGYA) are each inoculated with 1 g or 1 ml of product and the broths incubated at 308C. After incubation for 48 and 72 h, 0.1 ml from each broth is spread plated onto TGY agar containing 0.5% acetic acid and the plates incubated at 308C. The detection time of the method is shortened by incubating broths and plates at 308C rather than the traditional temperature of 258C, as the optimum growth temperature for Zygosaccharomyces bailii is 30–328C (Jermini and Schmidt-Lorenz, 1987b). The sensitivity of the method is greatly increased by using triplicate broths instead of single or duplicate broths and by spread plating 0.1 ml from each broth instead of streaking a loopful onto TGYA agar. This method has been used to detect low numbers of cells of Zygosaccharomyces bailii in experimentally inoculated cordial syrup, mayonnaise, salad dressing and barbecue sauce and other preservative-resistant yeasts such as Schizosaccharomyces pombe, Pichia membranaefaciens and some preservative resistant strains of Saccharomyces cerevisiae. Yeasts of intermediate preservative resistance (e.g. Debaryomyces hansenii, Candida krusei and Torulaspora delbrueckii) can also be detected by this method. Better recoveries were obtained using TGY than a malt extract agar and broth system, possibly due to the fact that the pH of TGY broth + 0.5% acetic acid is 3.8, compared with pH 3.2 for MEA + 0.5% acetic acid, and there is a lower concentration of glucose (2%) in the ME

4 Methods for Isolation, Enumeration and Identification

system compared with 10% in TGY. Yeasts which are unable to grow in the presence of acetic acid or other weak acid preservatives (sorbic or benzoic acids and their salts) are not detected by this method.

4.6.4 Techniques for Heat-Resistant Fungi Heat resistant spoilage fungi, such as Byssochlamys, Talaromyces, Neosartorya and Eupenicillium species can be selectively isolated from fruit juices, pulps and concentrates by laboratory pasteurisation using various methods (Beuchat and Rice, 1979; Hocking and Pitt, 1984; Beuchat and Pitt, 2001; Houbraken and Samson, 2006). Three methods are described here: a plating method based on that of Murdock and Hatcher (1978), a direct incubation method and a filtration method for liquid samples such as liquid sugar. Plating method. If the sample to be tested is more concentrated than 358Brix, it should first be diluted 1:1 with 0.1% peptone or similar diluent. For very acid juices such as passionfruit, normally about pH 2.0, the pH should be adjusted to 3.5–4.0. Two 50 ml samples are taken for examination. Erlenmeyer flasks (250 ml) or polyethylene Stomacher bags may be used as heat penetration into these containers will be rapid. If using Stomacher bags, the tops should be heat sealed. If a heat sealer is not available, the tops may be folded over and secured with a clip and should not be fully immersed. The two samples are heated in a closed water bath at 808C for 30 min, then rapidly cooled. Each 50 ml sample is then mixed with an equal volume of double strength MEA distributed over four 150 mm Petri dishes. The Petri dishes are loosely sealed in a plastic bag to prevent drying and incubated at 308C for up to 30 days. Plates are examined weekly for growth. Most moulds will produce visible colonies within 10 days, but incubation for up to 30 days will allow for the possible presence of badly heat damaged spores, which may germinate very slowly. This long incubation time also allows most moulds to mature and sporulate, aiding their identification. The main problem associated with this technique is the possibility of aerial contamination of the plates with common mould spores, which will give false positive results. The growth of green Penicillium

4.6 Choosing a Suitable Medium

colonies, or colonies of common Aspergillus species such as A. flavus and A. niger, is a clear indication of contamination as these fungi are not heat resistant. To minimise this problem, plates should be poured in clean, still air or a Class 2 biohazard cabinet if possible. If a product contains large numbers of heat resistant bacterial spores (e.g. Bacillus species), antibiotics can be added to the agar. The addition of chloramphenicol (100 mg/l of medium) will prevent the growth of these bacteria. Direct incubation method. A more direct method used for screening fruit pulps and other semisolid materials avoids the problems of aerial contamination. Place approximately 30 ml of pulp in each of three or more flat bottles such as 100 ml medicine flats. Heat the bottles in the upright position for 30 min at 808C and cool, as described previously. The bottles of pulp can then be incubated directly, without opening and without the addition of agar. They should be incubated flat, allowing as large a surface area as possible, for up to 30 days at 308C. Any mould colonies which develop will need to be subcultured onto a suitable medium for identification. If containers such as Roux bottles are available, larger samples can be examined by this technique, but heating times must be increased. Bottle contents should reach at least 758C for 20 min when checked by a thermometer suspended near the centre of the pulp. For further details of the above methods see Hocking and Pitt (1984) or Beuchat and Pitt (2001). Filtration method for liquid sugars. This method permits the detection of very low numbers of cells in clear liquids such as liquid sugar. Sample size should be at least 100 g, taken after vigorously shaking the container from which the sample is drawn. Add 100 ml diluent (0.1% aqueous peptone) to 2  50 g samples and mix well to dissolve. Filter both samples sequentially through the same sterile 0.45 mm membrane filter. After both samples have passed through the filter, rinse the interior of the funnel with 3  20– 30 ml volumes of sterile diluent. Remove the filter from the filter holder using sterile forceps and place it in a sterile bottle or Stomacher bag. Add 10 ml diluent to the bottle or bag containing the filter and place in a water bath at 758C for 30 min. Ensure the sample is submerged in the water bath (weigh down if necessary). Cool rapidly to room temperature, shake well, then divide the 10 ml of diluent between three Petri dishes. Add a generous portion of MEA with

33

antibiotics to each plate, mix the agar and sample well, then let the plates solidify. Incubate at 308C for up to 30 days, examining weekly. Count colonies and report count per 100 g. This method was developed by BCN Research Laboratories, Rockford, Tennessee, USA.

4.6.5 Other Plating Techniques Three other techniques which were developed for counting bacteria have been applied to fungal enumeration. Spiral plate count. Zipkes et al. (1981) evaluated the application of the spiral plate procedure to the enumeration of yeasts and moulds. They compared this procedure with the traditional pour plate and streak plate methods and found that spiral plating gave a higher overall recovery and lower replicate plating errors than the other two methods. The medium they used was potato dextrose agar, but the technique should be no less efficient using the media recommended here. Automation of spiral plate counting was studied by Manninen et al. (1991). They compared counts of pure bacterial, yeast and mould cultures using standard plating methods and spiral plate counts determined both manually and with a laser colony detector. They concluded that counts were not significantly different except where large colonies (10–15 mm diameter) of Rhizopus oligosporus were enumerated. Other Rhizopus and Mucor species are also likely to interfere with this method unless a suitable plating medium (such as DRBC) is used. Alonso-Calleja et al. (2002) found that the spiral plate technique compared well with standard plate counting on OGYE agar for enumeration of yeasts and moulds in goat’s milk cheeses; however, Garcı´ a-Armesto et al. (2002) found the method unsuitable for enumeration of yeasts in raw ewe’s milk. Hydrophobic grid membrane filters. Membrane filters overprinted with a square hydrophobic grid have been developed for rapid enumeration of bacteria. The hydrophobic grid membrane filter TM (HGMF) system is marketed as ISO-Grid by Neogen (Lansing, MI, USA). The HGMF ‘‘count’’ is determined by a most probable number (MPN) calculation. Brodsky et al. (1982) applied the HGMF technique to counting yeasts and moulds in foods. They compared it with spread plating on

34

potato dextrose agar amended with antibiotics and found that the HGMF technique produced higher counts in 2 days than the traditional method did after 5 days. The HGMF method could find use for evaluating the quality of raw materials before incorporation into a product. For example, Erickson (1993) has proposed a method using HGMF in conjunction with a selective medium for detection of the preservative resistant yeast Zygosaccharomyces bailii in acidified ingredients. The HGMF method was collaboratively trialled by 20 laboratories using 6 naturally contaminated foods. Although there were some differences between counts obtained by HGMF and the reference method (5 day pour plate), the differences were not significant, and the HGMF method was adopted by the AOAC International (Entis, 1996). Spangenberg and Ingham (2000) found that the HGMF method gave equivalent results to DRBC for enumeration of yeasts and moulds in grated low moisture Mozzarella cheese. Use of HGMF requires special holders for the square membrane filters, and an automated counting system is necessary to take full advantage of the method. The number of colonies on an HGMF can be counted visually, but it is a relatively slow process. TM Petrifilm Yeast and Mould. Petrifilm YM (3 M Company, St. Paul, MN, USA) is a proprietary system for enumerating fungi on a layer of medium enclosed in a plastic film, which eliminates the use of Petri dishes. A collaborative study carried out by the AOAC (Knight et al., 1997) concluded that Petrifilm YM gave comparable results with the BAM method (USFDA, 1992) for yeast and mould counts in five different food types inoculated with a cocktail of several species of Aspergillus and Penicillium and two species of yeast. Consequently, Petrifilm YM was adopted first action by AOAC International. Other workers (Beuchat et al., 1990, 2007; Taniwaki et al., 2001b; Ferrati et al., 2005) have shown that Petrifilm YM performed satisfactorily compared with conventional cultural methods when tested on a range of foods. However, Petrifilm was not particularly effective in inhibiting the growth of spreading moulds such as Rhizopus and Mucor. In addition, subculturing colonies for identification was more difficult from Petrifilm YM than from traditional Petri dishes. Petrifilm YM has a high aw and we do not recommend it for analysis of foods of less than 0.95 aw.

4 Methods for Isolation, Enumeration and Identification

4.7 Estimation of Fungal Biomass A deficiency in all of the enumeration techniques which rely on culturing fungi is that the result is at best poorly correlated with growth or biomass. Biomass is usually regarded as the fundamental measure of fungal growth in biotechnology, but it is not easy to quantify under the conditions existing in foods. Mycelial dry weight is most commonly used as a biomass estimate, but its relationship to mycelial wet weight and to metabolism varies widely in foods, due to the great influence of aw on both of the latter parameters. Fungi growing at reduced aw can be expected to be more dense than at high aw due to increased concentrations of internal solutes, though this is exceptionally difficult to measure experimentally. The question of a satisfactory fundamental measure of fungal biomass remains unanswered. Despite these basic problems, several chemical and biochemical techniques are available to estimate the extent of fungal growth in commodities. These techniques rely either on some unique component of the fungus that is not found in other microorganisms or foods or on immunological or molecular techniques. Some are still in the developmental phase: the most important ones are described briefly here.

4.7.1 Chitin Chitin is a polymer of N-acetyl-D-glucosamine and is a major constituent of the walls of fungal spores and mycelium. It also occurs in the exoskeleton of insects but is not present in bacteria or in foods. Hence the chitin content of a food or raw material can provide an estimate of fungal contamination. Chitin is most effectively assayed by the method of Ride and Drysdale (1972). Alkaline hydrolysis of the food sample at 1308C causes partial depolymerisation of chitin to produce chitosan. Treatment with nitrous acid then causes partial solubilisation and deamination of glucosamine residues to produce 2,5-anhydromannose, which is estimated colorimetrically using 3-methyl-2-benzothiazolone hydrazone hydrochloride as the principal reagent. Alkaline hydrolysis is more readily accomplished at

4.7 Estimation of Fungal Biomass

1218C in an autoclave (Jarvis, 1977). Improved assay sensitivity was achieved by derivatisation of glucosamine and other products with o-phthalaldehyde, separation by high performance liquid chromatography and detection of fluorescent compounds with a spectrofluorimeter (Lin and Cousin, 1985). Ekblad and Nasholm (1996) also described an HPLC method which measured fluorescence of a 9-fluorenylmethylchloroformate derivative of glucosamine. The chitin assay remains rather complex and slow, usually requiring about 5 h. A number of studies have indicated that the chitin assay is a valuable technique for estimating the extent of fungal invasion in foods such as maize and soybeans (Donald and Mirocha, 1977), wheat (Nandi, 1978) and barley (Whipps and Lewis, 1980) to estimate mycorrhizal fungi and fungal pathogens in plant material and soil (Ekblad and Nasholm, 1996; Ekblad et al., 1998; Penman et al., 2000; Singh, 2005) and measure wood rotting fungi (Nilsson and Bjurman, 1998). Particular attention has been paid to the possibility of developing the chitin assay as a replacement for the Howard mould count for tomato products (Jarvis, 1977; Bishop et al., 1982; Cousin et al., 1984). The chitin assay has some shortcomings and has been severely criticised by some authors (e.g. Sharma et al., 1977). The relationship between dry weight and chitin content varies at least twofold for different food spoilage fungi (Cousin et al., 1984; Lin and Cousin, 1985; Cousin, 1996). Some foods contain naturally occurring amino sugars such as glucosamine and galactosamine, which should be removed by acetone extraction prior to hydrolysis (Whipps and Lewis, 1980). Products from rot-free tomatoes give positive glucosamine assays even after acetone extraction (Cousin et al., 1984) and chitin content does not increase proportionally with fungal growth (Sharma et al., 1977). Insect contamination of grain samples has been reported to produce grossly misleading results (Sharma et al., 1977), but the presence of fruit flies in tomatobased products was less serious (Lin and Cousin, 1985). Materials such as stored grains frequently contain insect fragments and need to be checked before chitin assays are attempted. Because of these difficulties, the use of chitin as a chemical assay for fungi in foods has largely been superseded by the ergosterol assay.

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4.7.2 Ergosterol Ergosterol is the major sterol produced by fungi, but at most is a minor component of plant sterols (Weete, 1974). Ergosterol occurs as a component of fungal cell membranes, so is inherently likely to be correlated with hyphal growth and biomass. It is therefore a good candidate as a chemical for measuring fungal growth in foods and raw materials. Methodology for estimating ergosterol in cereals was developed by Seitz et al. (1977, 1979). Samples are blended with methanol, saponified with strong alkali, extracted with petroleum ether and fractionated by high pressure liquid chromatography. Ergosterol is detected by ultraviolet absorption, optimally at 282 nm, a wavelength at which other sterols exhibit little or no absorbance. For refinements to this methodology, see Newell et al. (1988). Liquid chromatography or gas chromatography with mass spectrometric detection (LC-MS; GC-MS) have also been used to estimate ergosterol (Headley et al., 2002; Dong et al., 2006; Varga et al., 2006). GC-MS in combination with nondiscriminating flash pyrolysis (Py-GC-MS) provides the advantages of little sample preparation and small sample size (Parsi and Gorecki, 2006). The ergosterol assay provides a useful method for quantifying fungal growth, so numerous studies have assessed the relationship between fungal growth and ergosterol production, using both liquid and solid substrates. Using liquid cultures, Zill et al. (1988) showed a correlation between ergosterol production, mycelial wet weight and mycelial protein in Fusarium graminearum. Matcham et al. (1985) reported that ergosterol correlated better with mycelial dry weight than chitin or laccase, a polyphenol oxidase. Variation exists, however. Torres et al. (1992) reported that Aspergillus ochraceus grown in liquid culture showed a threefold increase in ergosterol concentration in relation to mycelial dry weight as the culture aged from 2 to 26 days. Other reports (Matcham et al., 1985; Newell et al., 1987; Seitz et al., 1979) have indicated smaller variations, only one to twofold. Marfleet et al. (1991) showed that fungal biomass and ergosterol levels were correlated for three representative fungal species over a range of aw on solid substrates but not in liquid media. Nout et al. (1987) showed that the ergosterol content of Rhizopus oligosporus varied widely, from 2 to 24 mg/mg

36

mycelial mass, and varied with substrate, aeration and growth phase. The ergosterol content was low during the rapid growth phase but tended to increase, at times sharply, as growth slowed. Taniwaki (1995) demonstrated that in fungi growing in atmospheres low in O2 and with elevated CO2 levels, the ergosterol content of hyphae was significantly reduced. In atmospheres containing 60% CO2, ergosterol content per unit of hyphal length was up to six times less than in air. Growth medium also affected ergosterol concentrations: on average, seven foodborne fungal species grown on PDA produced more than twice as much ergosterol per unit hyphal length as when grown on CYA (Taniwaki, 1995; Taniwaki et al., 2006). However, there was a reasonable correlation between ergosterol and mycelium dry weight for seven of the eight species tested. Eurotium chevalieri was the exception: this species produced little ergosterol and appeared to produce several other sterols (Taniwaki et al., 2006). Marı´ n et al. (2005) examined 16 species of food spoilage fungi and concluded that ergosterol content and colony diameters were better correlated to fungal biomass than fungal counts were. Marı´ n et al. (2008) showed that, for 14 common food spoilage fungi, correlation coefficients between ergosterol and colony diameters were sufficiently significant over a range of aw values (0.95–0.85), pHs (5–7) and potassium sorbate concentrations (0.5–1.5%) for both parameters to be useful in growth modelling. Quantifying ergosterol production in foods has proved more difficult. Seitz et al. (1977) showed a good correlation between damage in rice grains and their ergosterol content, between ergosterol in wheat and rainfall during the growing season and between ergosterol content and fungal invasion in several sorghum hybrids. Matcham et al. (1985) reported good correlations between linear extension of Agaricus bisporus grown on rice grains and chitin, ergosterol and laccase production. Ergosterol content correlated with colony counts of fungi on wheat grains at 0.95 aw but not at 0.85 aw (Tothill et al., 1992). Using a stereomicroscope for visual examination, they concluded that sound grain contained up to 6 mg/g ergosterol, microscopically mouldy grain 7.5–10 mg/g and visibly mouldy grain more than 10 mg/g ergosterol. From studies on ergosterol levels, colony counts and mould growth in a variety of grain samples, Schnu¨rer and Jonsson (1992) concluded that ergosterol correlated with colony counts

4 Methods for Isolation, Enumeration and Identification

better on DG18 (r = 0.77) than on MEA (r = 0.69). Ergosterol levels of food grade wheat ranged from 2.4 to 2.8 mg/g dry weight, samples from field trials (of unspecified quality) from 3.0 to 5.6 mg/g and feed grains from 8 to 15 mg/g dry weight. After an extensive survey of ergosterol levels in Danish crops, Hansen and Pedersen (1991) concluded that the normal levels of ergosterol in barley were 7.6 – 2.8, wheat for bread making 5.0 – 1.5, rye for bread making 6.8 – 2.2, peas 2.2 – 2.7 and rapeseed 2.4 – 1.3 mg/g dry weight. Ochratoxin A in barley correlated well with ergosterol content and reached significant levels when ergosterol increased to 25 mg/g dry weight. However, aflatoxin B1 became detectable in cottonseed meal when ergosterol reached only 4 mg/g. ‘‘Burned’’ rapeseed, a measure of quality, became significant when ergosterol reached 1.4 mg/g dry weight. Lamper et al. (2000) found that ergosterol content correlated well (r = 0.87) with deoxynivalenol levels in wheat inoculated with Fusarium graminearum or F. culmorum. Moraes et al. (2003) found that there was good correlation between mould counts and ergosterol content of Brazilian maize (r = 0.94) but a poor correlation (r = 0.4) between ergosterol and aflatoxin content. Pietri et al. (2004) found significant correlation between ergosterol content of Italian maize and the major mycotoxins, fumonisin B1 (1995 crop) or zearalenone and deoxynivalenol (1996 crop). Ergosterol content correlated strongly with fat acidity values and germination ability of stored canola (Pronyk et al., 2006). These authors also noted that Penicillium and Aspergillus species contributed more to ergosterol than Eurotium species. Ergosterol levels in sound canola were between 1.46 and 1.67 mg/g, whereas levels above 2 mg/g indicated significant levels of spoilage (Pronyk et al., 2006). Karaca and Nas (2006) examined ergosterol content of dried figs and found good correlation (r = 0.92) between aflatoxin and ergosterol in reject figs which were fluorescent, but no significant correlation with patulin content. Kadakal et al. (2005) found good correlation (r = 0.98) between ergosterol and patulin in apple juice and that both patulin (r = 0.99) and ergosterol (r = 0.99) were linearly related to the proportion of decayed apples used to make the juice. Ergosterol has been used to assess mould growth in cheese with variable results (Pecchini, 1997; Taniwaki et al., 2001a).

4.7 Estimation of Fungal Biomass

Ergosterol content has also been investigated as an indicator of the mycological status of tomato products. Battilani et al. (1996) found a significant correlation between ergosterol, Howard mould count (HMC) and fungal growth, but with a high level of uncertainty. Kadakal et al. (2004) found a linear relationship between degree of decay in tomato pulp and HMC (r = 0.97) and ergosterol (r = 0.96) and concluded that ergosterol has the potential to be used in quality assessment of tomatoes. Sio et al. (2000) described an improved method for extraction of ergosterol from tomato products. Other applications of ergosterol as a measurement of fungal biomass include estimation of mould spores in indoor air and aerosols (Miller and Young, 1997; Robine et al., 2005; Lau et al., 2006), estimation of wood decay by fungi (Eikenes et al., 2005) and estimation of fungi in soil and wetlands (e.g. Headley et al., 2002; Zhao et al., 2005). The ergosterol assay is reported to have a high sensitivity and, in contrast to the chitin assay, requires only 1 h for completion (Seitz et al., 1979). Despite its limitations, it appears to be a useful indicator of fungal invasion of foods and to hold promise as a routine technique for quality control purposes.

4.7.3 Impedimetry and Conductimetry Metabolites produced by growth of microorganisms in liquid media alter the medium’s impedance and conductance. The use of changes in these properties as a measure of bacterial growth was suggested by Hadley and Senyk (1975) and was first applied to yeasts by Evans (1982) and to moulds by Jarvis et al. (1983). Most of the subsequent work on fungi has been carried out with yeasts, but the methodology is often applicable to moulds also. During the 1980s there were a number of studies aimed at optimising media for inducing detectable and reproducible changes in either conductance or capacitance during fungal growth (see Pitt and Hocking, 1997). Watson-Craik et al. (1989, 1990) studied 27 mould species on a wide range of both commercially available and specially prepared media and concluded that conductance and capacitance were both medium and species specific. The use of media high in ammonium ions and glucose, with added yeast extract and peptone, decreased the

37

influence of product variability and induced higher conductance changes (Owens et al., 1992). An impedimetric method for detection of heat resistant fungi in fruit juices was described by Nielsen (1992). The detection limit in artificially contaminated juices was one Neosartorya ascospore per millilitre, detectable in 100 h. Huang et al. (2003) reported an impedance method for detection of bacteria and fungi in bottled water which shortened the detection time from 5 days to 27.1 h for fungi and from 48 to 11.3 h for bacteria. Although impedimetry and conductimetry promised to be effective rapid methods when used under well-defined conditions for a specific purpose with a particular kind of food, the methodology does not appear to have been broadly taken up for food mycology applications.

4.7.4 Adenosine Triphosphate (ATP) ATP has also been suggested as a measure of microbial biomass because bioluminescence techniques provide a very sensitive assay (Jarvis et al., 1983). Provided that background levels of ATP in plant or other cells are very low, or that microorganisms can be effectively separated from such other materials, the method has some potential as a microbial assay. A good correlation was shown between ATP production and viable counts of six species of psychrotrophic yeasts grown in pure culture (Patel and Williams, 1985). The effective detection of low levels of yeasts in carbonated beverages by ATP has also been reported (LaRocco et al., 1985). However, living plant cells contain high levels of ATP and fungi are often very difficult to separate from food materials. Moreover, extraction of molecules from fungal cells is notoriously difficult, so this potential may be difficult to realise in food mycology. The most widespread application of ATP bioluminescence in the food industry is for monitoring hygiene of surfaces in food production facilities (Easter, 2007).

4.7.5 Fungal Volatiles Methods for detection and characterisation of fungal volatiles are finding increasing applications in

38

food mycology, for the detection of fungal deterioration of grain and other food products, for the identification of particular fungi and for assessing potential mycotoxin contamination. Over the last two decades, there has been rapid growth in the development of gas sensor arrays. Gas sensors are chemical sensors that produce an electronic signal which is used as input into a pattern recognition system in order to recognise different volatiles and odours. This integrated system of gas sensor and pattern recognition is often called an ‘‘electronic nose or e-nose’’ (Gardner and Bartlett, 1994; Schaller et al., 1998). Arrays comprising equipment to collect headspace volatiles, analysis by GC-MS or other sensitive analytical methods, coupled with computer analysis are also commonly used. Deterioration of stored grain results from a combination of chemical and biological changes, with changes due to fungal growth often predominant. Deterioration is marked by off-odour development, loss of germinability, caking, rancidity and sometimes mycotoxin development (Abramson et al., 1980). Fungi produce volatile chemicals during growth and particular chemicals may be associated with grain deterioration (Kaminski et al., 1974, 1975). Sinha et al. (1988) monitored production of 3-methyl-1-butanol, 1-octen-3-ol and 3-octanone in stored grain and found that the presence of these volatiles usually correlated with seed infection by Alternaria alternata, Eurotium repens, Aspergillus versicolor and several Penicillium species. Octenol and octanone in particular seemed to be associated with deterioration in grain due to fungal growth. Adamek et al. (1992) identified methylfuran, 2-methylpropanol and 3-methylbutanol as the most important metabolites from Eurotium amstelodami, Aspergillus flavus, Penicillium cyclopium and Fusarium culmorum growing on wheat. The volatiles produced by several Penicillium species were also studied. Borjesson et al. (1989) collected ¨ volatiles produced by several food spoilage fungi grown in pure culture on wheat grain and found that some compounds, especially 3-methyl-1-butanol, were produced early in fungal growth and could be used as an early warning of potential deterioration, before fungal growth became visible. The production of volatile metabolites by Penicillium species in grain correlated well with carbon dioxide production and ergosterol formation (Borjesson et al., 1990, 1992). The use ¨

4 Methods for Isolation, Enumeration and Identification

of volatile compound production as an indicator of mould deterioration in grains has been extensively assessed and reviewed (Kaminski and Wasowicz, 1991; Schnu¨rer et al., 1999; Magan and Evans, 2000; Paolesse et al., 2006; Balasubramanian et al., 2007). Fungal volatiles can be used to detect potential mycotoxin contamination, to discriminate between fungal species (Sunesson et al., 1995; Keshri et al., 1998) and even between toxigenic and non toxigenic strains of particular fungi (Sahgal et al., 2007). Karlshøj et al. (2007a) used an electronic nose to differentiate between closely related Penicillium species (P. camemberti, P. nordicum, P. paneum, P. carneum, P. roqueforti and P. expansum) from cheese. Volatile profiles can be used to predict mould spoilage in bakery products (Vinaixa et al., 2004; Marı´ n et al., 2007a) and to detect and differentiate between toxigenic and non toxigenic P. verrucosum strains in bakery products (Needham and Magan, 2003). Volatile profiles have also been used to differentiate between toxigenic and non toxigenic Fusarium strains (Keshri and Magan, 2000; Demyttenaere et al., 2004), to identify mycotoxins (aflatoxins, ochratoxin A and deoxynivalenol) in durum wheat (Tognon et al., 2005) and to detect and quantify ochratoxin A and deoxynivalenol in barley (Olsson et al., 2002). This technology has also been applied to predict the presence of P. expansum and patulin in apple products (Karlshøj et al., 2007b) and to detect and discriminate diseases of potato tubers (Kushalappa et al., 2002) and stemend rot and anthracnose in mangoes (Moalemiyan et al., 2006). Electronic nose technology has also been used for early detection of moulds in libraries and archives (Pinzari et al., 2004).

4.7.6 Immunological Techniques Cell wall proteins of fungi produce antigens, which can be detected by immunological methods. Some antigens are derived from components common to a wide range of fungi, and hence are indicative of general fungal growth, while others are genus or even species specific. A variety of methods have been developed to take advantage of these fungal antigens, although none is in widespread commercial use.

4.7 Estimation of Fungal Biomass

Enzyme-linked immunosorbent assay (ELISA). The preparation of antigens from three common foodborne fungi (Penicillium aurantiogriseum, Mucor racemosus and Fusarium oxysporum) was described by Notermans and Heuvelman (1985). Preparation of immunoglobulin antibodies against these antigens was followed by development of an ELISA assay. Fungi were detected in both unheated and heat processed foods by this method. Antigens were relatively genus specific: the M. racemosus antigens reacted with other Mucor and Rhizopus species, and the Penicillium antigen reacted with the Aspergillus species tested. It was subsequently shown that the Penicillium antigen reacted with 43 of 45 Penicillium species tested and that antigen production correlated with mycelial weight and was unaffected by culture conditions, medium, temperature and aw (Notermans et al., 1986). The Penicillium antigen also reacted with Aspergillus flavus and the level of antigen correlated with aflatoxin production (Notermans et al., 1986). ELISA techniques have also been studied as a potential replacement for the Howard mould count. Antigens from tomato moulds (Alternaria alternata, Geotrichum candidum and Rhizopus stolonifer) were used to produce an ELISA test sensitive to 1 mg/g of mould in tomato. A correlation was observed between antigen formation and mould added to tomato puree, while background interference was very low (Lin et al., 1986). The method was tested against a broader range of foods, with encouraging results (Lin and Cousin, 1987). Robertson and Patel (1989) improved the sensitivity of the method for tomato paste by using a polyclonal antibody against Botrytis cinerea, Mucor piriformis and Fusarium solani in addition to the three species used by Lin et al. (1986). ELISA based methodology has been reported for the detection of Fusarium species in corn (Meirelles et al., 2006), cornmeal (Iyer and Cousin, 2003) and grain (Rohde and Rabenstein, 2005). Correlations with other measures of fungal growth (ergosterol, colony count, mycotoxin levels) were variable. Aspergillus species have also been examined as targets for immunological detection because of their importance in mycotoxin contamination. ELISA detection of A. ochraceus in wheat (Lu et al., 1995), coffee powder, chilli powders and poultry feed (Anand and Rati, 2006) has been investigated, with reasonable correction with other parameters

39

(amount of ochratoxin A, r ¼ 0.93; percentage infection r ¼ 0.89) reported by Lu et al. (1995) but poorer correlation with colony forming units (r ¼ 68) and glucosamine content (r ¼ 0.64). Anand and Rati (2006) did not attempt to correlate their ELISA with external measurements of fungal biomass, but used an internal comparison method using freeze dried mycelium. Immunological detection of aflatoxigenic fungi has also been the subject of several studies, using antibodies raised to A. flavus or A. parasiticus (Candlish et al., 1997; Tsai and Yu, 1997, 1999; Yong and Cousin, 2001). Tsai and Yu (1999) reported good correlation between viable count and ELISA measurements, depending on substrate, with correlation coefficients varying from 0.96 for wheat to 0.86 for peanuts in artificially inoculated samples. ELISA tests for Botrytis and Monascus in foods (Cousin et al., 1990) and for the detection in rice of Penicillium islandicum (Dewey et al., 1990) and Humicola lanuginosa (Dewey et al., 1992) have been described. ELISA tests have also been applied to detection of yeasts in dairy products (Garcia et al., 2004) and fruit juice (Yoshida et al., 1996), moulds in paprika powder (Kisko et al., 1998) and moulds in agricultural commodities (Park et al., 2003) with varying success. Linfield et al. (1995) reported that polyclonal antibodies raised against Botrytis allii were useful in early detection of neck rot of onions caused by this pathogen. Immunochemical detection of fungi in food and feeds has been reviewed by Li et al. (2000). Latex agglutination. A different approach to the immunological detection of fungi in foods has been the coating of latex beads with antibodies and detection of agglutination of the beads in the presence of antigens (Kamphuis et al., 1989; Notermans and Kamphuis, 1992; Stynen et al., 1992). It was found that 0.8 mm latex beads coated with antibodies from the extracellular polysaccharide produced by Penicillium digitatum specifically detected Aspergillus and Penicillium species (Kamphuis et al., 1989). Detection limits were as low as 5–10 ng/ml of the purified antigen. Commercially produced latex agglutination tests were of value for screening the mycological quality of grains and processed foods (Braendlin and Cox, 1992; van der Horst et al., 1992), although one kit performed poorly in detection of mould in tomato products (van der Horst et al., 1992).

40

Schwabe et al. (1992) compared the latex agglutination assay with ergosterol production for detection of Penicillium, Aspergillus and Fusarium species in pure culture. They concluded that the two methods were comparable for Penicillium and Aspergillus but that ergosterol was more sensitive for Fusarium. In food samples, both the latex agglutination test and ergosterol were effective means of detecting mould growth, but no clear correlation existed in values obtained by the two methods. Kesari et al. (2004) used a latex agglutination test to detect teliospores of Karnal bunt (Tilletia indica) in single grains of wheat and were able to detect a few as 750 teliospores, which they reported as suitable for single seed analysis. Fluorescent antibody techniques. Fluorescent antibody techniques have also been used directly for the detection of mould in foods. Warnock (1971) detected Penicillium aurantiogriseum in barley by this method, while Robertson et al. (1988) used antisera from five fungi to visualise moulds and simplify their detection in the Howard mould count technique.

4.7.7 Molecular Methods The first techniques for detecting DNA sequences using specific probes were devised more than 30 years ago (Southern, 1975). With the development of the polymerase chain reaction (PCR), gene cloning and oligonucleotide synthesis, DNA sequences can now be prepared in large quantities for use in probes. Depending on its role in the genome, DNA may be specific at almost any taxonomic level. Molecular methods can be used in both detection and identification of fungi, and real-time PCR methods even allow quantitative detection. This field of research continues to develop very rapidly, and significant advances may be expected. Producing a broadly based or even strain specific DNA probe is theoretically possible for any organism. Full genome sequences are available for a number of important foodborne fungi, including several Aspergillus species (A. nidulans, A. fumigatus, A. oryzae, A. flavus, A. niger, A. terreus, A. clavatus and Neosartorya fischeri), Botrytis cinerea, Chaetomium globosum, Fusarium graminearum, F. oxysporum, F. verticillioides, Rhizopus oryzae and several yeast species (Saccharomyces

4 Methods for Isolation, Enumeration and Identification

cerevisiae, Candida albicans, C. guilliermondii, C. lusitaniae, C. tropicalis, Schizosaccharomyces japonicus and Sch. pombe) (Broad Institute, 2008). Penicillium species are noticeably absent from the list, although the genome of the pathogenic species P. marneffei has recently been sequenced (JCVI, 2008). There have been rapid advances in molecular methods for the detection of mycotoxigenic fungi. Probes have been developed that target genes in the mycotoxin biosynthetic or regulatory pathways for many species of Aspergillus, Penicillium and Fusarium, enabling these fungi to be detected and/or quantified in grains, grapes, apples, coffee and other food matrices (Edwards et al., 2002; Paterson, 2006; Geisen, 2007; Niessen, 2007a, b; Rossi et al., 2007). Molecular methods have also been developed for the detection of spoilage fungi, particularly yeasts, e.g. Brettanomyces/Dekkera in grapes and wine (Phister and Mills, 2003; Agnolucci et al., 2007; Hayashi et al., 2007), Hanseniaspora in wine (Phister et al., 2007); Zygosaccharomyces bailii in wine and fruit juices (Rawsthorne and Phister, 2006) and Kluyveromyces marxianus in yoghurt (Mayoral et al., 2006). More general techniques have been described for detection of yeasts in juices (Casey and Dobson, 2004; Ros-Chumillas et al., 2007), milk, yoghurt and cheese (Cocolin et al., 2002; Bleve et al., 2003; Garcı´ a et al., 2004; Lopandic et al., 2006; Gente et al., 2007), sourdough fermentation (Meroth et al., 2003), table olives (Arroyo-Lopez et al., 2006) and vacuum packaged ham (Sanz et al., 2005). PCR based methods have also been described for detection of moulds in orange juice (Wan et al., 2006), Alternaria in cereal grains (Zur et al., 2002) and Botrytis in onion seeds (Walcott et al., 2004). As gene sequencing has become more readily available and cheaper, along with freely accessible databases such as GenBank (http://www.ncbi.nlm.nih.gov/Genbank/), sequence analysis and comparison is becoming a routine method for identification of yeasts and filamentous fungi. For yeasts, the most common target region is the D1/D2 domain of the 26S rDNA (Kurtzman et al., 2003), but the internal transcribed spacer (ITS) region of ribosomal DNA (rDNA) may also be used (van der Vossen et al., 2003; Pulvirenti and Giudici, 2003; Solieri et al., 2006; Villa-Carvajal et al., 2006).

4.8 Identification Media and Methods

Sequencing of the ITS region along with ‘‘housekeeping genes’’ such as calmodulin, b-tubulin and elongation factor 1-a is now commonly applied for purposes of identification and phylogenetic analysis of important food spoilage and mycotoxigenic fungi in the genera Aspergillus (Varga, 2006; Geiser et al., 2007; Peterson, 2008), Penicillium (Peterson, 2004, 2006; Samson et al., 2004b; Wang and Zhuang, 2007; Serra et al., 2008) and Fusarium (O’Donnell et al., 2004; Scott and Chakraborty, 2006; Leslie et al., 2007). Despite the apparent power of molecular techniques, they need to be applied with some caution, particularly when comparing DNA sequences with those in the publicly available databases to identify yeast or mould isolates. Correct identification relies on the database sequences having the correct name attached to them by the depositor, which is not always the case. If the identification makes sense, the percent homology is 98% or greater and the number of base pairs on which the homology is scored is high, then the answer is probably correct.

4.8 Identification Media and Methods 4.8.1 Standard Methodology The identification keys in this book are based primarily on the standardised procedure described for the identification of Penicillium species by Pitt (1979b). Cultures are grown for 7 days on three standard media at 258C, and on one of these at 5 and 378C also. The three Media are Czapek yeast extract agar (CYA; Pitt, 1973) used at all three temperatures; malt extract agar (MEA; Raper and Thom, 1949) and 25% glycerol nitrate agar (G25N; Pitt, 1973). Their formulae are given in the Media Appendix. Preparation time of CYA and G25N is reduced by the use of Czapek concentrate (Pitt, 1973), which is added to the media at the rate of 1% of the aqueous portion. As media ingredients have become more purified in recent years, difficulties with extent and colour of sporulation on CYA have been encountered, especially with some Penicillium species. To overcome this problem, Czapek concentrate has been reformulated (Pitt, 2000) by the inclusion of traces of zinc and copper (Smith, 1949) (see Media Appendix).

41

4.8.2 Plating Regimen As noted above, cultures for identification are grown on three media and at three temperatures. Maximum efficiency in time, incubator space and materials is achieved by inoculating two cultures on a single Petri dish of G25N, and at 5 and 378C, as shown in Fig. 4.3. Cultures are rarely mutually inhibitory under these conditions, although overgrowth of one culture by another is occasionally a problem at 378C. Standard sized Petri dishes (90–100 mm) are used, except that at 58C, smaller plates (50–60 mm in diameter) can be an advantage. The smaller size is easier to examine under the low power microscope. All plates are incubated for a standard time of 7 days. Plates incubated at 378C should be enclosed in polyethylene bags to prevent evaporation and drying of the medium. Unless the humidity is very low, plates at 258C will not dry excessively in 7 days. If no 58C incubator is available, use a polyethylene food container or insulated box in a household refrigerator. The box should be equipped with a thermometer, and its location moved by trial and error until a place with a 58C average temperature is located. Temperatures at 5 and 378C should ideally be –0.58C and be checked frequently; at 258C, –28C control is adequate (Okuda, 1994).

4.8.3 Inoculation As shown in Fig. 4.3, Petri dishes of CYA and MEA for incubation at 258C are inoculated with a single culture at three points, equidistant from the centre and the edge of the plate and from each other. Plates of the other media are inoculated with two points per culture, as illustrated. With some fungi, especially Penicillium and Aspergillus, it is important to minimise colonies from stray spores. The most satisfactory technique is to inoculate plates with spores suspended in semisolid agar (Pitt, 1979b). Dispense 0.2–0.4 ml of melted agar (0.2%) and detergent (0.05%), such as polysorbitan 80 (Tween 80), in small vials and sterilise. To use, add a needle point of spores and mycelium to a vial and mix slightly. Then, before flaming the needle, use it to stab inoculate the 58C plate;

42

4 Methods for Isolation, Enumeration and Identification

Fig. 4.3 Schematic of regimen used for culturing fungal isolates for identification

residual spores on the needle make a good inoculum. Next, take a sterile loop, mix the vial contents thoroughly and inoculate the standard plates. Used vials can be sterilised by steaming and reused several times before being washed or discarded.

4.8.4 Additional Media and Methods The plating regimen outlined above can be used to identify most of the fungi described in subsequent chapters of this book. Some exceptions exist, because certain genera either grow poorly or fail to sporulate on the standard media. As noted earlier in this chapter, fastidious xerophiles are identified on MY50G agar. Eurotium species, traditionally identified on Czapek agar with 20% sucrose, are identified here on Czapek yeast extract agar with 20% sucrose (CY20S; see Appendix). Trichoderma species are best identified on potato dextrose agar (PDA) after a relatively short incubation time (3–4 days), as the structures tend to autolyse as cultures mature. Penicillium subgenus Penicillium. Many species classified in subgenus Penicillium are morphologically similar, and identification using traditional morphological techniques remains difficult. These

Penicillia are very common in foods, and many produce mycotoxins, so correct identification is often critical. Species within subgenus Penicillium fall into two groups: those with an affinity for proteinaceous foods and those which grow more vigorously in foods high in carbohydrate. Frisvad (1981, 1985) introduced creatine sucrose agar (CREA) to permit differentiation between these two groups. Creatine (as the sole nitrogen source) permitted growth of the former group while inhibiting the latter. Incorporation of bromocresol purple enabled visualisation of pH changes, either acid or alkaline, depending on the creatine or sucrose metabolism of a particular species. However, discrimination between positive and negative responses was not always clear, and the main tabulation of species reactions to CREA (Table 1 in Frisvad, 1985) was difficult to interpret. Frisvad (1993) subsequently produced a number of variations of CREA, including acid and neutral pH formulations and substitution of sucrose with fructose or lactose. Although the merits of each formulation were discussed, no firm recommendation resulted from this exercise. Pitt (1993a) modified Frisvad’s strongly alkaline CREA medium by studying several sucrose and creatine concentrations over a wide pH range. The result was neutral creatine sucrose agar (CSN), a medium producing eight different reactions among

4.8 Identification Media and Methods

the 20 Penicillium subgenus Penicillium species tested. When included in the normal plating regimen for identification of Penicillium cultures from the subgenus Penicillium, CSN provides a very useful aid to distinguishing between these difficult and closely related species. See Chapter 7 for details of use of CSN, its reactions and interpretation. Its formulation is given in the Media Appendix. Dematiaceous Hyphomycetes. The natural habitats for many dematiaceous Hyphomycetes are plants or plant material, so finding suitable laboratory media and conditions to induce typical sporulation can be difficult. Alternaria, Curvularia, Stemphylium and Ulocladium species are best identified from dichloran chloramphenicol malt extract agar (DCMA; Andrews, 1992b) plates incubated for 7 days at 258C under lights. Conidial characteristics of Bipolaris species vary with type of medium. Species described here are best identified from tap water agar (TWA) containing a natural substrate such as sterilised wheat or millet seeds, or wheat straw. Drechslera species will grow well on DCMA but sporulation is poor. The best sporulation is achieved with V-8 juice (V-8 J) agar or TWA with one of the natural substrates mentioned above, incubated at 258C for 7–10 days under lights with a 12 h photoperiod. Light/dark periodicity is important as Drechslera cultures require light to produce conidiophores and darkness to produce conidia, possibly due to light inactivation of flavine necessary for conidial formation (Knan, 1971; Platt et al., 1977). For the identification of Trichoderma and Fusarium species, potato dextrose agar (PDA) is used. Fusarium species also require additional methods and media as outlined below.

4.8.5 Identification of Fusarium Species Fusarium isolates exhibit unusually high variability in colony morphology and also may deteriorate rapidly in culture. Thus, they should be identified as soon as possible after initial isolation with a minimum of subculturing to avoid deterioration. It is common practice to prepare cultures of Fusaria from single spores for growth on identification media, as this reduces both of these problems.

43

Single sporing. The technique for preparing single spore cultures is as follows (Nelson et al., 1983; Leslie and Summerell, 2006). Pour about 10 ml of 2% water agar into unscratched glass or plastic Petri dishes and allow to dry, either by holding the plates at room temperature for several days or by placing them inverted in an oven at 37–458C for about 30 min. Prepare a suspension of conidia in a 10 ml sterile water blank so that it contains 1–10 spores per low-power (10) microscope field when a drop from a 3 mm loop is examined on a slide. With experience, this concentration can be gauged simply by observing the turbidity of the suspension. Pour the suspension of spores onto a dried water agar plate, drain off the excess liquid and incubate in an inclined position at 20–258C for 18–20 h. After incubation, open the Petri dish, shake off any accumulated moisture droplets and examine under a stereomicroscope using transmitted light. The germinating conidia should be visible under 25 magnification. A dissecting needle with a flattened end and sharpened edges is used to cut out small squares of agar containing single, germinating conidia. These single conidia are then transferred on the agar blocks to the desired growth medium. If the original culture is contaminated with bacteria, a drop of 25% lactic acid may be added to the water blank. Allow this acidified spore suspension to stand for 10 min before pouring onto a water agar plate. Germination of acid-treated Fusarium conidia may be delayed by 24 h or more. Media. Two media have been used in this book for the identification of Fusarium isolates: potato dextrose agar (PDA) for colony characteristics and colour and dichloran chloramphenicol peptone agar (DCPA) for the development of diagnostic macro-, micro- and chlamydoconidia. A third medium, carnation leaf agar (CLA), is recommended by some Fusarium specialists for Fusarium cultivation and identification (Nelson et al., 1983; Leslie and Summerell, 2006). CLA is an excellent medium, on which most Fusarium species readily produce their diagnostic macroconidia. Production of macroconidia on DCPA is usually comparable with that on CLA, but microconidia and chlamydoconidia are often more plentiful on CLA due to greater production of aerial hyphae. DCPA is used in the present work rather than CLA,

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however, because dried, gamma-irradiated carnation leaves are difficult to obtain in many localities. Inoculation and incubation. For identification by the methods used in this book, single spore cultures of Fusarium isolates should be prepared on agar blocks as outlined above, inoculated, one per plate, onto two plates each of PDA and DCPA and incubated at 258C for 7 days. Individual plates may be used for each medium, or alternatively divided plates may be used, with one medium on each half of the plate. Illumination during incubation is essential for the production of macroconidia. The light source may be diffuse daylight (not direct sunlight) or light from a bank of fluorescent tubes. A photoperiod of 12 h per day is normally used. Alternating temperatures of 20 and 258C have been recommended (Nelson et al., 1983; Leslie and Summerell, 2006) but are not essential. A simple light bank may be constructed from a standard 40 watt fluorescent fixture with two cool white tubes, suspended 0.5–l m above the laboratory bench or shelf supporting the cultures. The addition of a black light tube (e.g. Philips TL 40 W/80 RS F40BLB) is also desirable and in some cases essential to induce macroconidial or chlamydoconidial production.

4.8.6 Yeasts Yeast identification systems. Identification of foodborne yeasts remains a difficult task, as colony characteristics and microscopic morphology are of limited value. Generally it has been necessary to use biochemical and physiological tests such as fermentation of carbohydrates, assimilation patterns for a range of carbon and nitrogen sources and growth at various temperatures. Details of these methods and media may be found in Kurtzman and Fell (1998), Kurtzman et al. (2003) or Barnett et al. (2000). Molecular methods (see below) are being used increasingly in yeast identification, replacing time consuming biochemical testing. Identification using systems based on biochemical and physiological testing is complex and time consuming. However, a number of attempts have been made to assist those who wish to persevere with yeast identification. Several simplified systems

4 Methods for Isolation, Enumeration and Identification

have been published in the literature, and both automated and manual yeast identification systems are now commercially available. Dea´k and Beuchat (1987) published a simplified identification key (SIK) which included 215 species of foodborne yeasts. They subsequently modified their system, restricting it to the 76 species most frequently occurring in foods (SIM), and reported that it was much more successful than the API 20C system (BioMe´rieux, Marcy-l’Etoile, France) for identification (Dea´k, 1992). The SIM uses only two Petri dishes and three test tubes to examine each strain for ability to assimilate 10 carbon sources, fermentation of glucose, assimilation of nitrate and splitting of urea. These biochemical tests are supplemented by morphological observations. SIM separates the yeasts into six groups by a dichotomous key utilising the results of five key tests. Further tests are used to differentiate the yeasts in these six groups using secondary dichotomous keys. A dichotomous key to 25 common species of foodborne yeasts was published by Smith and Yarrow (1995), who used 17 biochemical and physiological tests to distinguish the species. Of the commercial systems available, the most widely used for foodborne yeasts are the Biolog, which is an automated system, and the BioMe´rieux ID32C yeast identification strips which can be read manually or automatically using the ATB system. BioMe´rieux also markets the fully automated VITEK 2 system, based on a card containing 64 tests. However, the database comprises only 46 clinically important species (Aubertine et al., 2006) and so has limited application to foodborne yeasts. The Biolog (Biolog Inc., Hayward, CA, USA; http://www.biolog.com/microID.html) utilises a yeast identification test panel (YT MicroPlateTM) consisting of a matrix of 8  12 wells. The first three rows contain 35 carbon source oxidation tests using tetrazolium violet as an indicator of oxidation. The next five rows contain carbon assimilation tests which are scored turbidimetrically against a negative control panel containing only water. The last row contains two carbon sources and tests for the co-utilisation of various carbon sources with D-xylose. The hardware (Biolog MicroStation Reader) consists of an automated plate reader coupled with a computer, which interprets the

4.9 Examination of Cultures

results and compares them with the resident database which currently includes 267 species. Manual interpretation of the Biolog plates is not recommended. This system has been designed with the food industry in mind, and the database contains all the common foodborne yeasts, unlike other systems which are usually aimed at the clinical market. The ATB ID32C system (BioMe´rieux, Marcyl’Etoile, France) is an automated system utilising the BioMe´rieux ID32C yeast identification strips. These strips contain 30 assimilation tests plus a positive (glucose) and a negative control well, all of which are inoculated with a yeast suspension of specified density. The strips are incubated at 308C for 48–72 h. As with the Biolog, the ATB automated system consists of a plate reader attached to a computer. The database associated with the ATB system contains 63 yeast species. The BioMe´rieux ID32C strips can be read manually, and the results enable identification of yeasts using published keys or computer identification programmes such as that of Barnett et al. (1996). This system may be used with reasonable success, particularly when the test results from the ID32C strips are supplemented with extra tests (glucose fermentation, urease production, nitrate utilisation, growth in 0.5 and 1% acetic acid and in 50 and 60% glucose, growth at 378C, production of pseudohyphae, ascospore formation and morphological observations) to give a more comprehensive base for the identification. Growth in 0.5 and 1% acetic acid indicates preservative resistance, and growth in 50 and 60% glucose gives an indication of ability to grow at reduced aw. Both these parameters are important in determining a yeast’s ability to cause spoilage in particular products. Identification using DNA sequencing is increasingly becoming the method of choice, as extensive databases such as GenBank are freely available for identification purposes. The 600– 650 nucleotide D1/D2 region of the large subunit (26S) ribosomal DNA is the most widely targeted section of the genome, and sometimes the ITS region may also be used (Kurtzman et al., 2003). Sequencing of the D1/D2 region, along with some supplementary physiological and biochemical tests, is the identification method currently used in our laboratory.

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Even using the available systems, identification of yeasts still requires some specialist knowledge and interpretation and remains time consuming. Our experience indicates that no more than 12 species of spoilage yeasts are of real concern in foods. It is possible to differentiate these few species by relatively simple techniques, i.e. colony and microscopic morphology, growth on the standard media used for filamentous fungi, growth on other media which test for preservative resistance, ability to use nitrate as a nitrogen source and adaptation to high NaCl concentrations. Details of these techniques are given in Chapter 10.

4.9 Examination of Cultures As noted above, all cultures for identification should first be grown on the standard regimen described earlier. After 7 days incubation, the following examination should be carried out and then the general key to fungi in Chapter 5 should be used. That key will be of assistance even in the event of cultures failing to grow under one or more of the standard conditions.

4.9.1 Colony Diameters Measure the diameters of macroscopic colonies in millimetres from the reverse side (Fig. 4.4). Microscopic growth or germination at 58C is assessed by low power microscopy (60–100), by putting the 58C Petri dish on the microscope stage and examining by bright field transmitted light. Growth at 378C is assessed macroscopically only; germination of spores at 378C is an unreliable character.

4.9.2 Colony Characters Colony appearance can be judged by eye or with a hand lens, but examination is more effective if a stereomicroscope is used. Magnifications in the range of 5–25 are the most useful. Characters such as type and location of sporing structures and

46

4 Methods for Isolation, Enumeration and Identification

Fig. 4.4 Technique for measuring colony diameters by transmitted light

extent of sporulation are best gauged with the stereomicroscope. Reflected light is usually more effective than transmitted light. To determine colony colours, examine colonies by daylight or by daylight-type fluorescent light. In some genera, reference to a colour dictionary is helpful. The Methuen ‘‘Handbook of Colour’’ (Kornerup and Wanscher, 1978) has been used in this work and is highly recommended.

4.9.3 Preparation of Wet Mounts for Microscopy Fungi should always be examined microscopically as wet mounts rather than fixed and stained like bacteria. To prepare a wet mount, use an inoculating needle to cut out a small portion of the colony which includes sporing structures. Examination with the stereomicroscope can be an invaluable aid here. For freely sporing fungi with little mycelium, cut a piece of colony near the edge, where fruiting structures are young and spore numbers not excessive. Take structures which may enclose spores, i.e. cleistothecia, from near colony centres, where the probability of mature spores is highest. If the only differentiated parts of the colony appear to be

buried in the agar, e.g. pycnidia, take a sample of these with a small piece of the agar. Float the cut colony sample from the needle onto a slide with the aid of a drop of 70% ethanol. It may be necessary to tease out the specimen with the needle and the corner of a cover slip (square cover slips are best). Fungal specimens may be highly hydrophobic: the ethanol helps to wet the preparation, minimising the amount of entrapped air. When most of the ethanol has evaporated, add a drop of lactic acid (for phase or interference contrast optics) or lactofuchsin stain (see below) for bright field. Add a cover slip; if necessary remove excess liquid from the preparation by gently blotting with facial tissue or similar absorbent paper. The preparation is now ready for examination.

4.9.4 Staining A wide variety of stains are in use for mycological work. However, most are time consuming to prepare, or to use, or are slow to act, because fungal walls and spores are highly resistant to stains. By far the most effective stain for use in food mycology is lactofuchsin (Carmichael, 1955), which suffers from none of these faults. It consists of 0.1% acid fuchsin

4.9 Examination of Cultures

dissolved in lactic acid of 85% or higher purity. Young actively growing fungal structures are preferentially coloured bright pink, so sporing structures can usually be readily distinguished against a background of older mycelium. Cleistothecial initials, developing asci and maturing ascospores are also seen more readily in preparations stained with lactofuchsin. Like most other mycological stains, lactofuchsin is corrosive. Take care to clean it off microscope parts or skin! Be especially careful of the objective faces, because lactic acid will slowly corrode the relatively soft glass used in lenses.

4.9.5 Microscopes and Microscopy Choice of microscope. Fungal identifications inevitably involve microscopy. A high quality, binocular compound microscope is essential for serious mycological work. Bright field, phase contrast and interference contrast optics are all suitable. If bright field optics are used, preparations should be stained. Phase contrast optics avoid the need for staining, although as explained above, staining has merit for certain structures in any case. Resolution under phase contrast sometimes suffers from excessive halo effects because fungal structures are highly refractile. Experience suggests that bright field optics are more satisfactory than phase contrast for examining most fungi. Interference contrast is superior to both bright field and phase contrast and is strongly recommended. The very thin optical section cut by this system provides higher resolution of dense structures and the relatively low contrast is restful to use. The microscope should be equipped with at least four objectives: 6 for examining Petri dishes under low power (for germination, etc.); 10 or 16 for searching fields for sporing structures; 40 for examining structural details and measuring stipes, fruiting structures and large conidia; and 100 oil immersion for observation of details of spore attachment, surface texture, ornamentation of hyphae and spores and fine measurements. Oculars should be 10 or 12.5 and may with advantage be wide field and have a high eye point suitable for use with spectacles. One should be a focusing ocular,

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equipped with an eyepiece micrometer, which is essential for measuring dimensions of spores and sporogenous structures. In the examination of fungal mounts, it is stressed that it is most important to use low power optics before succumbing to the temptation to use oil immersion. The principal reason is that fungal preparations usually remain as small clumps and do not disperse as bacteria do. Only under low power is the search for the optimal area of the slide for the observation of fruiting structures likely to be rewarded. Once a suitable area is located under the 10 or 16 objective, move to the 40. This should be the lens most used; microscope optics are such that only the finest details of ornamentation can be observed more effectively under oil immersion than at this magnification. Aligning the microscope. Correct alignment of the microscope is essential, so that its resolution is as high as possible and it can be used for long periods without discomfort. An incorrectly aligned microscope will lead to poor observation, discomfort, fatigue, headache and eye strain. A person of normal visual acuity should be able to use a correctly aligned instrument throughout a whole working day without discomfort. The steps to correctly align a microscope are given below. They should be read in conjunction with the microscope manufacturer’s instructions. 1. Mount a slide on the stage and bring it into approximate focus. If a prepared slide is not available, a slide marked with a marking pen or ink line is a satisfactory substitute. 2. Close the microscope’s field diaphragm (the one at the microscope base nearer the light source). The image of the diaphragm opening should now be visible in the microscope field. If it is, first focus it with the condenser focusing knob and then centre it in the field with the condenser centring screws. If the diaphragm opening cannot be seen, first rack the condenser up and down and watch to see if the opening becomes visible; if it does not, rack the condenser to its highest position and then slowly open the field diaphragm until the opening comes into view. Centre the diaphragm approximately and proceed as above. 3. For bright field optics, the condenser diaphragm should be adjusted each time the objective power is changed. Remove one ocular; close the

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condenser diaphragm so that the field seen down the open tube is about two-thirds its maximum size. With phase contrast and interference contrast systems, this adjustment is less critical. The preceding steps align the microscope itself and should be checked frequently. If optimal illumination is desired, each step should be carried out for each new slide and each objective change. As a routine habit, the whole process should take only a few seconds. If the available microscope is not equipped with a built-in light source, a field diaphragm and a fully centring condenser, it is unlikely to be satisfactory for the identification of small spored fungi such as Penicillium species. The following steps are designed to align the observer with the microscope, compensating for individual differences in sight. Provided settings on the microscope are remembered, these steps need be carried out only occasionally, to check that visual acuity has not altered. Different settings will be needed for an individual with and without spectacles or contact lenses. 4. Assuming the microscope is binocular, pull the oculars out to their greatest distance apart and then, while watching a focused field, move them gradually together until a single circular field is seen without strain or head movement. Note the distance on the scale between the oculars; this is the individual’s interpupillary distance. Repeat this operation two or three times until satisfied that the correct distance has been found. This distance should always remain the same and be similar on any microscope. 5. Under the 40 or 100 objective, locate a tiny, readily recognised point on the slide and focus on it. Take a piece of white card and place it between the focusing ocular and the corresponding eye. Leave the eye open. Now focus the tiny point with the other eye, carefully, with the microscope fine focus. Next, transfer the white card to the other ocular and, using the focusing collar beneath the ocular, refocus the tiny point. Remove the card and note the setting on the scale. Repeat until satisfied the correct setting has been found. 6. On some microscopes, the eyepiece micrometer can be focused independently. Use the focusing system on the ocular itself to focus the micrometer. Note the setting on the scale on the side of the ocular.

4 Methods for Isolation, Enumeration and Identification

Always check the settings on the microscope before use and after making measurements with the micrometer. It is very easy to upset the ocular alignment when measuring.

4.10 Preservation of Fungi Many fungi are stable in culture and can be subcultured repeatedly without apparent change or deterioration. Others, especially Fusarium species and other plant pathogens and some mycotoxigenic species, will degenerate rapidly after only a few transfers. For stable isolates that are used routinely in the laboratory, storage on agar slopes is satisfactory. Many freely sporing fungi will survive for several months, and sometimes much longer, when stored at 1–48C on a medium such as CYA. One of the hazards of storage at these temperatures is contamination by psychrophilic Penicillium or Cladosporium species. Storage at freezer temperatures (–18 to –208C) prevents such contamination, but some fungi do not survive well under these conditions. Storage at room temperatures, at 108C or above, for long periods is not advisable because of the likelihood of invasion by culture mites (see below).

4.10.1 Lyophilisation For unstable cultures, and indeed for the long term storage of any food spoilage fungi, freeze drying or lyophilisation is probably the best method of preservation. Many commercial systems are now available for carrying out this process. A satisfactory menstruum for lyophilisation of most fungi is l.5 normal strength reconstituted nonfat milk powder (15% in distilled water). For fungi with hydrophobic conidia, such as Aspergillus and Penicillium, a small amount of detergent (0.05%) such as polysorbitan 80 (Tween 80) should be added to the milk. Dispense the milk in 10 ml lots in small tubes or 12.5 ml (0.5 oz) McCartney bottles, and sterilise by steaming for 20 min on three successive days (the Tyndallisation process). The milk must be stored at 208C or above between steamings, to permit bacterial spores to germinate.

4.10 Preservation of Fungi

Occasionally bacterial spores will survive this process: it is advisable to store the milk at room temperature for some days after Tyndallisation. Any bottles which show clotting or other breakdown should be autoclaved and discarded. Autoclaving at 1218C, even for 10 min or less, is not recommended, as browning will occur and browning compounds are known to be inhibitory to microorganisms. Most common spoilage fungi survive lyophilisation well. However, in our experience and that of others (Mikata et al., 1983; Smith and Onions, 1994), some yeasts, plant pathogens and xerophiles do not. Storage of lyophilised ampoules at refrigeration temperatures (0–48C) is recommended, but room temperature is probably satisfactory provided sunlight is avoided and temperatures do not exceed 308C. Some laboratories routinely store lyophilised cultures at –18 to –208C. Strains being maintained for a particular trait or utilised for a specific purpose such as system testing or metabolite production should always be lyophilised. Continued subculturing often leads to deterioration or loss of the desired character. The ability of a strain to produce a particular mycotoxin, for example, may decrease with each transfer. Isolates should be lyophilised as soon as possible after primary isolation to prevent degeneration.

4.10.2 Other Storage Techniques A variety of systems other than lyophilisation have been proposed for long term storage of fungi. Of these, liquid nitrogen storage has found most acceptance with major culture collections. This type of storage appears to be superior to any other for plant pathogens and fungi which will not sporulate in pure culture. However, liquid nitrogen systems are expensive to establish and maintain and are only suitable for large collections. Freezer units which run at very low temperatures (–808C or below) are available and are well suited to the needs of culture collections. In our laboratory we routinely store cultures at –808C, using glycerol (60–80%) as a cryoprotectant. Spore suspensions are prepared by taking conidia or ascospores from a freely sporulating sector of the colony, dispersing them in the glycerol then freezing in screw capped

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cryovials. An 80% solution of glycerol remains viscous at –808C, which enables cultures to be removed from the freezer for subculture without the need for defrosting. For smaller laboratories that do not have access to lyophilisation, liquid nitrogen or ultra-low temperature storage, some simple techniques exist that can be used to maintain fungal cultures over relatively long periods (e.g. 1–10 years) without the need for subculturing. Water storage. A simple and inexpensive method of fungal culture preservation is storage of agar blocks in water (Smith and Onions, 1994). Small agar blocks (7–10 mm2) are cut from the growing margin of a young fungal colony and placed in sterile water in a bottle such as a Bijou bottle (6.25 ml or 0.25 oz McCartney bottle). The rubber lined cap is screwed down and the bottles stored in a cool room (1–108C). Cultures may be revived by removal of a block and placing it on a suitable growth medium. Using this method, cultures have been reported to remain viable and retain their characteristics for up to 7 years (Boeswinkel, 1976; Smith and Onions, 1994). Some yeasts may be maintained by storing as suspensions in water (Kirsop, 1984). Growth from a late logarithmic slant culture is suspended in sterile distilled water and transferred to a sterile container so that 90% of the volume is filled with the suspension. Containers are stored at room temperature. Survival of some Candida, Saccharomyces, Cryptococcus, Rhodotorula and Schizosaccharomyces species for up to 4 years has been reported (Kirsop, 1984). Silica gel storage. Many fungal cultures may be maintained for long periods (often more than 10 years) by drying spore suspensions onto silica gel (Smith and Onions, 1983; Smith, 1984). This method is not suitable for mycelial cultures, but can be used with some success for yeasts (Kirsop, 1984). As silica gel liberates heat when moistened, the technique depends on keeping the cultures cool enough to avoid damaging the spores during preparation. Medium grain plain (non-indicating) silica gel of 6–22 mesh is placed in suitable glass bottles (Bijou or McCartney bottles) to a depth of about 1 cm and sterilised, either by dry heat at 1808C for 2–3 h or by autoclaving at 1218C for 15 min. Autoclaved silica gel must be thoroughly

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dried in an oven before use. Bottles are precooled by placing in a tray of ice or refrigerating for 24 h, then transferring to an ice tray for inoculation. Suspensions of fungal spores or yeasts are prepared in sterile skimmed milk (as for lyophilisation, above) and the suspension added to the cooled silica gel to wet three-quarters of it. The bottles and gel are allowed to dry at room temperature for 10–14 days with caps slightly loosened. Caps are then screwed down and the bottles stored at 48C (storage at room temperature is also satisfactory) in air-tight containers over indicating silica gel to absorb any moisture. Cultures are revived by shaking a few crystals of silica gel onto a suitable growth medium (broth culture may be better for yeasts). Survival varies according to species or even strain, but survival of yeasts for up to 5 years (Kirsop, 1984) and fungi for more than 10 years (Smith and Onions, 1983) has been reported.

4.11 Housekeeping in the Mycological Laboratory Like any other microbiological laboratory, a mycological laboratory should be kept in a clean condition. Discard unwanted cultures regularly and dispose of them by steaming or autoclaving. Wipe bench tops regularly with ethanol (70–95%). Floors should be wet mopped or polished only with machines equipped with efficient vacuum cleaners and dust filters. Where possible store food and plant materials away from the laboratory. Open Petri dishes carefully. Use small inocula on wet needles. Transport Petri dishes to the stereomicroscope stage before removing lids. Do not bump cultures during transport. Contrary to popular belief, a well-run mycological laboratory is not a source of contamination to bacteriological laboratories. The air in a mycological laboratory should not carry a significant population of fungal spores. The reverse problem can occur, however, because bacteria multiply more rapidly than do fungi. Bacterial spores are often present in food laboratories, readily infect fungal plates and can rapidly outgrow and inhibit fungal mycelia, especially at 378C.

4 Methods for Isolation, Enumeration and Identification

If for any reason fungal spore concentrations do build up in a laboratory and cause an unacceptable level of contamination, the air should be purified. The simplest technique is to spray the air throughout the laboratory with an aerosol before it is closed in the evening. Any aerosol spray, such as a room deodoriser or air freshener, is effective. Aerosol droplets entrain fungal spores very efficiently and carry them to the floor. A more drastic and effective treatment in cases of severe contamination is to spray a solution of 2% thymol in ethanol around the room and close it for a weekend. The spray is rather pungent, and while not harmful to humans, it effectively kills fungal spores and mites (see below). Do not leave cultures on benches before fumigation.

4.11.1 Culture Mites A major hazard in growing and maintaining fungal cultures is the culture mite. Many species of mites live on fungal hyphae as their main or sole diet in nature and find culture collections an idyllic environment. Mites crawl from culture to culture, contaminating them with fungi and bacteria as they go or, given long enough, may eat them out entirely. Mites are very small (0.05–0.15 mm long), usually just visible to the observant naked eye. They are arachnoids, related to spiders, and hermaphroditic. Each mite leaves a trail of eggs about half adult size as it goes. Eggs hatch within 24 h and reach adulthood within 2 or 3 days. The damage an unchecked mite plague can do to a fungal culture collection has to be experienced to be believed. The most common sources of mites are plant material, soil, contaminated fungal cultures and mouldy foodstuffs. Mites can also be carried on large dust particles. Building work near a laboratory almost always induces a mite infestation. The avoidance of losses due to mites requires constant vigilance. Always watch for telltale signs, such as contaminants growing around the edges of a Petri dish, a ‘‘moth-eaten’’ appearance to colonies or ‘‘tracks’’ of bacterial colonies across agar. Examination of suspect material or cultures under the stereomicroscope will readily reveal the presence of mites and mite eggs.

4.11 Housekeeping in the Mycological Laboratory

Adult mites are rapidly killed by freezing, and mite eggs will only survive 48–72 h at –208C. Cultures contaminated by mites can often be recovered by freezing for 48 h, then subculturing from uninfected portions of the culture with the aid of the stereomicroscope. Suspect food and other samples being brought into the laboratory should also be frozen for 24 h to destroy mites before enumeration or subculturing is carried out. Infestation by mites can be minimised by good housekeeping, i.e. by avoiding accumulation of dust or old cultures in the laboratory. It is also good practice to handle and store food and plant samples well away from areas where fungi are inoculated and incubated. To control a mite infestation, remove all contaminated material, including cultures. Freeze Petri dishes and culture tubes which must be recovered; autoclave, steam or add alcohol to all others. Clean benches thoroughly with sodium hypochlorite (household bleach) or 70% ethanol. Incubators can be disinfested with aerosol insecticides.

4.11.2 Problem Fungi There are three fungal invaders which should be watched for carefully in a food mycology laboratory: Aspergillus fumigatus, Rhizopus stolonifer and Chrysonilia sitophila. The first is a human pathogen; the others can cause a contamination chain which is difficult to break. Aspergillus fumigatus may cause invasive aspergillosis in the lungs or serious allergenic responses in some individuals. It is sound practice to immediately kill and discard cultures of this fungus as soon as it is recognised. On no account should it be used for experimental studies in food spoilage or biodeterioration without precautions to prevent dissemination of spores. The morphology of A. fumigatus is described in detail in Chapter 8, but it is readily recognisable in the unopened Petri dish:

 colonies are low, dull blue and broadly spreading, with a velvety texture;

 growth is very rapid at 378C, covering a Petri dish in 2 days;

 long columns of blue conidia are readily seen under the stereomicroscope.

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Rhizopus stolonifer is a ubiquitous fungus in many kinds of foods. It grows rampantly at 258C, filling a Petri dish with sparse, dark mycelium in 2 days. It produces barely macroscopic aerial fruiting structures which are at first white, then become black. Given seven undisturbed days, it sheds dry, black spores outside the Petri dish, providing an effective inoculum for a continuous chain of future contamination. Once such an infection occurs, it is essential to carefully place the contaminated plate in a suitable container such as a plastic bag before transporting it to steamer or autoclave. Then use 70% ethanol to clean areas on which the plate had been incubated or placed. At daily intervals, carefully examine all plates inoculated subsequently, discarding any which show Rhizopus contamination, until infection ceases. Spraying the air with aerosols or thymol in ethanol will assist. Unlike A. fumigatus, R. stolonifer is not pathogenic. The third problem fungus, Chrysanilia sitophila, is known as ‘‘the red bread mould’’. It used to be common, but due to changes in manufacturing practice, it is now encountered less frequently, either in the bakery or in the laboratory. Like Rhizopus stolonifer, it grows with great rapidity at 258C. It forms a thin, pink mycelial growth across a Petri dish, clearly following the oxygen gradient which leads to the open air. It will force its way unerringly between dish and lid and, once outside, will produce masses of pink spores, which are quickly shed and build up around the base of the Petri dish. Decontamination relies on the same techniques as for Rhizopus: C. sitophila is probably the more difficult fungus to eradicate. Again, apart from its nuisance value in the laboratory, it is harmless.

4.11.3 Pathogens and Laboratory Safety While it must be said that any fungus which is capable of growth at 378C is a potential mammalian pathogen, the physiology of the healthy human is highly resistant to nearly all of the fungi encountered in the food laboratory. Nevertheless, fungi which can grow at 378C should be treated with caution. In particular, the habit of sniffing cultures is to be avoided wherever possible. It is true that odours produced by fungi have been used quite frequently as taxonomic criteria, especially in older

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publications, but their subjective and ephemeral nature makes them of little value for this purpose, and the risks involved are serious. Some laboratories regard fungal volatiles as such a serious risk that cultures such as Penicillium and Aspergillus species are handled in a biohazard cabinet. Many types of fungal spores are allergenic or carry mycotoxins. Inhalation of fungal spores should be avoided as far as possible. Of the fungi described in this book, only the Aspergilli normally pose any direct threat to health. Aspergillus fumigatus has already been mentioned, and care should also be taken when handling cultures of Neosartorya species, which are closely related to

4 Methods for Isolation, Enumeration and Identification

A. fumigatus and also grow prolifically at 378C. Other Aspergillus species, particularly A. flavus, A. niger and A. terreus, have been isolated from human specimens from time to time. For more details, see de Hoog et al. (2000). These species appear to be mainly opportunists and pose little threat to healthy people. Careful handling and good housekeeping are all that are required. However, immunocompromised individuals are in a different category. It is increasingly evident that such people have little resistance to fungal infection. Persons suspected to be immunocompromised, regardless of the cause, should not work in a mycology laboratory nor indeed be permitted knowingly to enter one.

Chapter 5

Primary Keys and Miscellaneous Fungi

Principles underlying fungal classification have been outlined in Chapter 3, including a brief overview of the relevant divisions of the Kingdom Fungi and their principal methods of reproduction. Some further detailed information is necessary in this chapter to assist in the use of the keys which follow. Ascomycetes. As discussed in Chapter 3, Ascomycetes produce ascospores in asci (Fig. 3.2). One genus, Byssochlamys, produces asci which are unenclosed; all other genera of relevance here produce asci in some kind of fruiting body, or ascocarp. The two kinds of ascocarp commonly seen in food spoilage fungi, the cleistothecium and the gymnothecium, have been described and illustrated in Chapter 3 (Fig. 3.3). Both types of ascocarp are usually pale or brightly coloured, not dark, and release ascospores by rupturing irregularly. Of genera relevant here, cleistothecia are produced by Emericella, Eurotium, Eupenicillium, Monascus and Neosartorya, and gymnothecia by Talaromyces. A third class of ascocarp, less commonly encountered in foodborne fungi, is the perithecium. Perithecia have cellular walls like cleistothecia, but are distinguished by the presence of an apical pore or ostiole through which asci or ascospores are liberated; also asci are long and clavate with ascospores arranged linearly within them. In the one perithecial genus of interest here, Chaetomium, the perithecia are black and have stout hyphae attached to the walls (Fig. 5.10). Conidial fungi. The strictly conidial fungi, also known as Fungi imperfecti or Deuteromycetes or anamorphic fungi, possess an amazing variety of ways of producing conidia. Terminology for structures bearing conidia and for conidia themselves has

become astonishingly complex in recent years; fortunately most of it is not essential for the recognition of the genera discussed in this text. Terms which are important in the keys which follow are described below. A fundamental division within the asexual fungi separates genera which form conidia aerially, grouped in the class Hyphomycetes, from those in which conidia are borne in some sort of enveloping body, the class Coelomycetes. Hyphomycetes. Fungi have developed seemingly endless ways of extruding or cutting off conidia, solitarily or in chains, from fertile cells which themselves may be borne solitarily or aggregated into more or less ordered structures. Hyphomycete taxonomy attempts to thread a way through this maze. In general, type and degree of aggregation of the fertile cells, and type of conidium, provides the basis for generic classification, while details of these characters and of spore size, shape and ornamentation are more commonly used to distinguish species. Features of conidia used in the keys in this work are length, septation, ornamentation and colour, particularly whether walls are light or dark. The method of conidium formation (ontogeny) is seldom emphasised here, because terminology is complex and distinctions may not be obvious. The principal point to note is the disposition of conidia: they may be borne solitarily, i.e. just one conidium per point of production; singly, i.e. successively from a single point, but unattached to each other; or in chains. Solitary conidia are borne on a relatively broad base and usually adhere to the fertile cell. Conidia formed in chains are usually extruded from a small cell of determinate length, often a

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_5, Ó Springer ScienceþBusiness Media, LLC 2009

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phialide, which in most genera narrows to a distinct neck. Conidia borne singly may be extruded in this same manner, or be borne by extrusion from a pore in a hypha or fertile cell, or be cut off by hyphal fragmentation. Phialidic Hyphomycetes. Hyphomycetes may produce phialides solitarily (the genus Acremonium) or in loosely ordered structures (Trichoderma) or highly ordered structures (Aspergillus, Penicillium and related genera). Genera of interest here with less ordered phialidic structures can mostly be differentiated by macroscopic characters, e.g. colony diameters and colours. However, differentiating genera with highly ordered phialidic structures will necessitate careful microscopic examination. Phialides in Aspergillus, Penicillium and related genera are sometimes borne directly on a stalk or stipe which arises from a hypha; in other species, however, the phialides are borne from supporting cells, termed metulae (sing. metula) and in some species the metulae may in turn be supported by other cells, termed branches or rami (sing. ramus). The whole structure, including the stipe, is called a conidiophore. In Aspergillus, stipes are always robust, with thick walls and usually without septa; the stipe terminates in a more or less spherical swelling, the vesicle, which bears phialides, or metulae and phialides, over most of its surface. In Aspergillus, phialides (and metulae) are always produced simultaneously, and this feature can readily be recognised by examining young developing conidiophores (Fig. 8.1a). Similar structures, though smaller, are produced by some Penicillium species: these are clearly distinguished from Aspergillus species by stipes which are septate and by phialides which are produced over a period of time (successively; Fig. 8.2b). Most Penicillium species, and those of related genera, do not produce phialides on vesicles, but in a cluster directly on a stipe, or on metulae and/or rami. The fruiting structure in Penicillium and related genera is termed a penicillus, while that in Aspergillus (for want of a better term) is called a head (or more recently, an aspergillum, Klich, 2002). Coelomycetes. As noted earlier, Coelomycetes produce conidia within an enveloping body, termed a conidioma (pl. conidiomata). In Petri dish culture, conidiomata are produced on or just under the agar surface and are macroscopically visible, usually being 100–500 mm in diameter. Two kinds of conidioma are important here: the pycnidium, a more

5 Primary Keys and Miscellaneous Fungi

or less spherical body with one or more pores (ostioles) through which conidia are released, and the acervulus, a flat body from which conidia are released by lifting or rupturing of a lid. The majority of Coelomycetes are pathogens on plants and many have not been studied in pure culture. In consequence, their taxonomy is difficult and genera and species are often poorly delimited. For a complete account of Coelomycete taxonomy see Sutton (1980). Yeasts. Yeasts are fungi which have developed the ability to reproduce by forming single vegetative cells by budding or, in a few species, by fission, in a manner similar to bacteria. Like bacteria, and unlike fungal spores, such cells are metabolically active and may in turn reproduce by budding (or fission). Yeast cells may survive for long periods both in culture and in nature. In consequence many yeasts produce true spores rarely or not at all. Yeasts are readily distinguished from filamentous fungi on the agar plate by their soft-textured colonies and limited growth. They are usually also readily distinguished from bacteria by their raised and often hemispherical colonies, white or pink colours and lack of ‘‘bacterial’’ odour. If in doubt, make a simple wet mount of a colony in water or lactofuchsin, add a cover slip and examine with the oil immersion lens. Yeast cells are larger than bacteria, measuring at least 3  2 mm and are nonuniform in size. If the culture is not too old, some cells will usually show developing buds. Yeasts cannot be classified solely by morphological features or growth on the standard media, and so are considered in a separate chapter (Chapter 10).

5.1 The General Key The taxonomic terms discussed above will enable the use of the general and miscellaneous keys which follow, although some other taxonomic terms may be introduced in discussions of particular genera. It is emphasised that these keys are designed for use on isolates which have been incubated for 7 days on the standard plating regimen outlined in Chapter 4. Colony diameters should be measured in millimetres from the reverse side by transmitted light. The general key has been designed to be as simple as possible and suitable for routine use, but it should be read in conjunction with the notes below it.

5.1 The General Key

55

General key to food spoilage fungi 1

No growth on any standard medium in 7 days Growth on one or more standard media

Chapter 9 – ‘‘Xerophilic fungi’’ 2

2 (1)

Colonies yeasts, either recognisably so on isolation or in culture, i.e. colonies soft, not exceeding 10 mm diam on any standard medium Growth filamentous, exceeding 10 mm diam on one or more standard media

Chapter 10 – ‘‘Yeasts’’ 3

3 (2)

Growth on CYA and/or MEA faster than on G25N Growth on G25N faster than on CYA and MEA

4 (3)

Hyphae frequently and conspicuously septate Hyphae lacking septa or septa rare

5 (4)

No mature spores present in 7 days Mature spores present in 7 days

6 9

6 (5)

Immature fruiting structures of some kind present No fruiting structures (or spores) detectable by lowpower microscopy or wet mounts from CYA or MEA

7

5 Chapter 6 – ‘‘Zygomycetes’’

7 (6)

Colonies and fruiting structures white or brightly coloured Colonies or fruiting structures dark

8 (7)

Colonies and fruiting structures white Colonies or fruiting structures brightly coloured

9 (5)

Spores (conidia) less than 10 mm long, borne in chains on clustered fertile cells (phialides), on well-defined stipes Spores (conidia) of various sizes, borne singly or solitarily, or if borne in chains, then chains not in aggregates

10 (9)

11 (10)

Conidia blue or green, phialides produced successively on vesicles, vesicles less than 10 mm diam, stipes usually septate Conidia variously coloured, phialides and/or metulae produced simultaneously on vesicles, vesicles larger than 10 mm diam, stipes nonseptate

Couplet 1. No growth on any standard medium indicates an extreme xerophile, i.e. Xeromyces bisporus or a Chrysosporium species, or a nonviable culture. Next inoculate culture onto MY50G for 7 days at 258C. If growth occurs, enter the key in Chapter 9, ‘‘Xerophilic Fungi’’; no growth on MY50G indicates a nonviable culture. Chrysosporium and Xeromyces

See section on ‘‘Miscellaneous fungi’’ below 8 Continue incubation; when spores mature, refer to section on ‘‘Miscellaneous fungi’’ below Chapter 8 – ‘‘Aspergillus and its teleomorphs’’ Chapter 7 – ‘‘Penicillium and related genera’’

Phialides or metulae and phialides borne on more or less spherical swellings on the stipe apices Phialides borne on penicilli, i.e. on unswollen stipes with or without intervening metulae and rami

5.1.1 Notes on the General Key

4 Chapter 9 – ‘‘Xerophilic fungi’’

10 See section on ‘‘Miscellaneous fungi’’ below

11 Chapter 7 – ‘‘Penicillium and related genera’’

Chapter 7 – ‘‘Penicillium and related genera’’

Chapter 8 – ‘‘Aspergillus and its teleomorphs’’

isolates are usually white or rarely golden brown. If the original culture used as inoculum is coloured other than pure white or golden brown, it is probably nonviable. Couplet 2. Yeasts are usually readily distinguished by slow growth, soft, easily sampled colonies, small spherical to ellipsoidal cells, often of variable size and shape and by reproduction by budding. See Chapter 10, ‘‘Yeasts’’ for identification procedures.

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5 Primary Keys and Miscellaneous Fungi

Couplet 3. The ability to grow more rapidly on G25N than on CYA or MEA indicates a xerophile (at least for keying purposes here). Check the key in Chapter 9, ‘‘Xerophilic Fungi’’. Some isolates can be identified from the standard plates, while others will require growth on CY20S or MY50G for identification. Couplet 4. The absence of septa in young, growing hyphae indicates an isolate belongs to subkingdom Zygomycotina, discussed here in Chapter 6 ‘‘Zygomycetes’’. Couplets 5, 6. Some isolates from a variety of fungal genera will not produce spores on the standard media in 7 days. Continue to incubate such cultures, preferably in diffuse daylight such as a laboratory window sill, at temperatures near 258C. Also inoculate such cultures onto two or three plates of DCMA and incubate these at 258C or thereabouts in darkness and in diffuse daylight or, if possible, under fluorescent illumination (see Chapter 4). After 1–2 weeks, check again for spores or fruiting bodies. If such structures are not seen, the isolate is unlikely to be significant in foods. Apparently asporogenous cultures should also be checked with a stereomicroscope while scraping up a sector of the colony with a needle. Fruiting bodies submerged in the agar will sometimes become visible with this technique. Couplet 7. Some isolates which produce white or brightly coloured fruiting bodies also produce very sparse aerial conidial structures which are easily overlooked. Check such cultures carefully with the stereomicroscope; if conidial structures are found, make a wet mount and reenter the key at Couplet 5.

A finely drawn glass needle will sometimes be of assistance in making mounts from delicate conidial structures on the colony. Nearly all dark fruiting structures encountered will mature at 258C within 2 weeks. Light does not usually influence this process. When mature spores are formed, refer to the following section.

5.2 Miscellaneous Fungi In this section are considered the genera which do not logically fit into some larger grouping considered elsewhere. Some are important in specific food spoilage problems, others are found in particular habitats such as cereals, while still others represent the aerially dispersed fungal flora found as ubiquitous contaminants or saprophytes. As will be seen, they are a very heterogeneous collection. Most fungi significant in food spoilage or food contamination and not treated in other chapters are included here. It is inevitable, though, that occasional isolates from foods will not belong to the genera considered in this section. The key has not been designed to take account of this, as it would be a practical impossibility. So when an isolate appears to key out satisfactorily, it must be checked against the description to confirm the identification. Some isolates will of course belong to a recognisable genus, but not the species described; in that case the references indicated will provide further information. The miscellaneous fungal genera are considered in alphabetical order following the key.

Key to miscellaneous genera 1

Colonies on CYA and MEA not exceeding 60 mm diam in 7 days Colonies on CYA or MEA exceeding 60 mm diam in 7 days

2 12

2 (1)

Conidia borne within a fruiting body on or beneath the agar surface Conidia borne from aerial or surface hyphae

3 (2)

Mycelium and conidia hyaline or brightly coloured Mycelium and/or conidia dark coloured

4 11

4 (3)

Conidia with a single lateral septum Conidia nonseptate or with more than one septum

Trichothecium 5

5 (4)

Conidia borne from gradually tapering fertile cells (phialides) Conidia borne directly on hyphae or by budding or hyphal fragmentation

Phoma 3

6 7

5.2 Miscellaneous Fungi

6 (5)

Colonies exceeding 50 mm diam on CYA Colonies not exceeding 50 mm diam on CYA

7 (5)

Colonies exceeding 45 mm diam on MEA; conidia borne solely by the breakup of hyphae to form arthroconidia Colonies not exceeding 40 mm diam on MEA; conidia not exclusively arthroconidia

8 (7)

9 (8)

Conidia exceeding 12 mm long; developing cleistothecia, fist-like on arm-like stalks, also present Conidia not exceeding 12 mm long; no evidence of cleistothecial development Conidia yeast-like, borne on spicules (small projections) from hyphae, or by budding; conidia 5 mm or less long Some conidia yeast-like, borne by budding; hyphal fragments, arthroconidia and/or chlamydoconidia also present; conidia >5 mm long

10 (9)

Budding hyphal fragments (10–50 mm long) present Hyphal fragments absent; or if present, not budding

11 (3)

Colonies low, mucoid and yeast-like, becoming grey to black in both obverse and reverse Colonies dry and velutinous, obverse green, reverse olive or deep blue black

57

Fusarium Acremonium

Geotrichum 8

Monascus 9

Endomyces 10 Hyphopichia Moniliella

Aureobasidium Cladosporium

12 (1)

Spores borne within an enclosed fruiting body on or under the agar surface Spores borne from aerial or surface hyphae

13 17

13 (12)

Spores consistently less than 15 mm long The larger or all spores more than 15 mm long

14 15

14 (13)

Fruiting bodies perithecia with stout, black hyphae attached to the walls Fruiting bodies pycnidia, without attached hyphae

15 (13)

Fruiting bodies roughly spherical (pycnidia) Fruiting bodies flat (acervuli)

16 (15)

Conidia hyaline or brightly coloured, nonseptate, without terminal appendages Conidia dark, with three or four septa and with spike-like, sometimes branched, terminal appendages

Chaetomium Phoma Lasiodiplodia 16 Colletotrichum Pestalotiopsis

17 (12)

Colonies and conidia hyaline or brightly coloured Colonies and/or conidia dark coloured

18 22

18 (17)

Colonies with grey or green areas Colonies white, orange, pink or purple

19 20

19 (18)

Colonies green Colonies grey

20 (18)

Colonies low and persistently white Colonies floccose, white or becoming brightly coloured

Geotrichum 21

21 (20)

Colonies predominantly orange, orange conidia shed profusely around the Petri dish rim Colonies white, pink or purple, sporulation on MEA weak or absent, better on DCPA under lights

Chrysonilia

22 (17)

Conidia consistently less than 15 mm long Conidia frequently exceeding 15 mm long

Trichoderma Botrytis

Fusarium 23 25

58

23 (22)

5 Primary Keys and Miscellaneous Fungi

Conidiophores long, branched, apically swollen, bearing closely packed pale brown conidia Conidiophores short or ill-defined, dark brown or black conidia borne irregularly

Botrytis 24

24 (23)

Conidia dark brown, often with a lighter coloured band around the periphery Conidia uniformly jet black

Arthrinium Nigrospora

25 (22)

Conidia approximately spherical Conidia elongate

Epicoccum 26

26 (25)

Conidia with transverse septa (or thick walls between cells) only Larger conidia with both transverse and longitudinal septa or irregularly septate

27 30

27 (26)

Conidia clavate (club shaped), often with long hyphal appendages at the apices; found almost exclusively on rice Conidia cylindroidal, ellipsoidal or an elongate ‘‘D’’ shape; source more general

Trichoconiella 28

28 (27)

Conidia cylindroidal, with parallel sides except at the terminal cells Conidia fusoid, narrowing from the central cells to the terminal cells, often bent or ‘‘D’’ shaped

Drechslera

29 (28)

Conidia not exceeding 40 mm long Conidia usually exceeding 40 mm long

Curvularia Bipolaris

30 (26)

Conidia clavate (club shaped) Conidia spherical to roughly ellipsoidal or short cylindrical

Alternaria 31

31 (30)

Conidia often tapering towards the base and sometimes pointed, i.e. pyriform or apiculate Conidia spherical to short cylindrical, not tapering from base to apex, with rounded ends

5.3 Genus Acremonium Link Commonly referred to as Cephalosporium Corda in pre-1970 literature, Acremonium is a large and varied genus characterised by the production of small, hyaline, single-celled conidia borne singly, i.e. successively but not connected to each other, from solitary phialides. A variety of species have been recorded from foods from time to time. One species, A. strictum, described here, is of relatively common occurrence. In this species, the phialides gradually taper to the apices without basal thickening or formation of a distinct neck, and conidia aggregate in balls of slime. Under the stereomicroscope, the slime balls look like large single spores, but their true nature becomes evident in wet mounts. A. strictum appears mainly in the earlier food literature under the name C. acremonium. As noted by Domsch et al. (1980), this name has been used for a variety of species, so that reports on physiology and occurrence are unreliable (Fig. 5.1).

Acremonium strictum W. Gams

29

Ulocladium Stemphylium

Fig. 5.1

Cephalosporium acremonium (name of uncertain application; no valid authority).

Colonies on CYA 20–30 mm diam, white or orange to pink, dense to floccose or funiculose; reverse pale or with orange to pink tones. Colonies on MEA 13–20 mm diam, similar to those on CYA or of slimy texture. Colonies on G25N <5 mm diam, usually 1–2 mm, of white mycelium. Sometimes growth at 58C. No growth at 378C. Conidia borne successively but singly from the apices of long, solitary, usually unbranched phialides, aggregating in a slime ball at the phialide tip, cylindrical to ellipsoidal, hyaline, 3–6  1–2 mm, smooth walled. Distinctive features. See genus preamble. Taxonomy. As noted above, Acremonium strictum is the correct name for the fungus commonly called C. acremonium in the earlier food literature (Gams, 1971).

5.3 Genus Acremonium

59

Fig. 5.1 Acremonium strictum (a) colonies on CYA and MEA, 7 d, 258C ; (b, c) phialides, bar = 10 mm; (d) conidia, bar = 5 mm

Physiology. Mycelial growth and sporulation of Acremonium strictum were found to be enhanced by the addition of mannitol to MEA (Teixeira et al., 2005). Mycotoxins. A. strictum and other Acremonium species encountered in foods are not known to produce mycotoxins. Ecology. The main source of Acremonium strictum is maize (Pitt et al., 1993; Pitt and Hocking, 1997; Williams et al., 1992). It has recently been

isolated from black bean seeds in Argentina (Castillo et al., 2004) and raw cork (Quercus suber) in Spain (Alvarez-Rodriguez et al., 2002). Other records, including those of Acremonium spp. or the earlier name Cephalosporium acremonium, include wheat, barley and rice, bananas showing crown rot, fresh vegetables, peanuts, pecans, hazelnuts and walnuts, soybeans, frozen meat, salami and biltong (Pitt and Hocking, 1997). References. Gams (1971); Domsch et al. (1980).

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5 Primary Keys and Miscellaneous Fungi

5.4 Genus Alternaria Nees: Fr. Alternaria produces large brown conidia with both longitudinal and transverse septa, borne from inconspicuous conidiophores, and with a distinct conical narrowing or ‘‘beak’’ at the apical end. These conidia are often formed in chains. Two other genera, Stemphylium Wallroth and Ulocladium Preuss, produce similarly septate conidia, but are uncommon in foods. Those of Ulocladium are roughly ellipsoidal, usually somewhat narrower at the base, and at most have small beaks. Both Alternaria and Ulocladium produce new conidia by lateral growth of the conidiophore from near the apex or through the beak area. Stemphylium conidia look like those of Ulocladium, but new conidia never form from a beak. New conidia are produced through the pore from which a conidium has already been produced. For a more detailed account of the distinctions among these three genera, see Simmons (1967).

Trichoconiella forms conidia similar to those of Alternaria, but with transverse septa only. This genus is associated with rice. Emory Simmons has spent a lifetime working with Alternaria and related genera and has recently revised the taxonomy of Alternaria comprehensively (Simmons, 2007). The simple notions of earlier authors, i.e. that Alternaria alternata is everywhere and that specific plants were colonised by specific pathogens, have been painstakingly replaced by a much more robust taxonomy. For example, Alternaria from citrus fruits may be Alternaria citri, but may also be one of 20 other species. The monograph by Simmons (2007) includes 276 species and is an essential laboratory manual for anyone who wishes to identify Alternaria species from a range of sources. Alternaria is not an easy genus for the inexperienced, however. Treatment here is confined to three common species only, with cereals as their major source.

Key to included species 1. Conidia produced in unbranched chains on DCMA Conidia produced in branched clusters on DCMA

Alternaria tenuissima (see A. alternata) 2

2. Conidia in unbroken chains, i.e. narrow hyphal elements between spores lacking; species occurring on wheat and many other substrates Primary conidia producing secondary conidia, i.e. conidia often separated by short hyphal lengths; species usually occurring on wheat

Alternaria alternata (Fr.) Keissl.

Fig. 5.2

Alternaria tenuis Nees

Colonies on CYA and MEA 50–70 mm diam, or covering the whole Petri dish, plane, of deeply floccose off-white to grey brown mycelium; reverse brown to nearly black. Colonies on G25N 10–15 mm diam, low and dense, olive brown or grey; reverse brown to almost black. At 58C, at least microcolonies, often colonies up to 4 mm diam. Usually no growth at 378C, occasionally colonies up to 10 mm diam, similar in appearance to those at 258C, or white. Colonies on DCMA 60–70 mm diam or covering the whole Petri dish, plane, floccose but not sparse, mycelium grey to dark grey; reverse grey to dark grey or black, less commonly bluish black. Conidia blown out from the apices of undistinguished conidiophores as short, irregularly branched

A. alternata A. infectoria

chains of up to 10 units, and then septating both laterally and longitudinally, with up to six transverse and two to three longitudinal or oblique septa, usually of clavate or pyriform shape overall, tapering towards the apices, forming a short beak, in culture usually 20–40  8–12 mm, with walls smooth to conspicuously roughened. Distinctive features. A. alternata is the most common saprophytic Alternaria species, occurring on a wide variety of sources, in contrast to other species considered here. On DCMA, conidia are produced more or less uniformly above the agar surface, in quite lengthy chains. The larger conidia have both longitudinal and transverse septa. Taxonomy. The first edition of this book noted that with acceptance of the 1821 starting point date for Alternaria nomenclature, A. alternata slowly became accepted as the correct name for this

5.4 Genus Alternaria

61

a

b

c

Fig. 5.2 Alternaria alternata (a) colonies on PDA and DCMA, 7 d, 258C ; (b) conidia in situ, bar ¼ 50 mm; (c) conidia, bar ¼ 20 mm

species. With the reversion to be the 1753 starting point date, the correct name again became A. tenuis. However, publications by Fries, 1821–1832, were later given special status under the ICBN, so A. alternata is once more the correct name. Further changes appear unlikely. Physiology. Optimum growth of Alternaria alternata is near 258C with minima variously reported as – 5 to 6.58C and maxima near 368C (Hasija, 1970; Domsch et al., 1980). The minimum aw for growth at 258C is 0.86 (Hocking et al., 1994). Optimal growth occurs at pH 4–5.4, and the pH range for growth is 2.7–8.0 (Hasija, 1970). A. alternata is able to grow in oxygen concentrations as low as 0.25%

(v/v) in N2, with growth rates being proportional to oxygen concentration (Follstad, 1966; Wells and Uota, 1970). The presence of the volatile compounds 3-methyl-1-butanol, 1-octen-3-ol and 3octanone in stored wheat grains is usually correlated with infection by A. alternata, amongst other fungi (Sinha et al., 1988). Mycotoxins. Alternaria alternata produces several toxins, of which the most important is tenuazonic acid (TA) (Logrieco et al., 2003). Tenuzoic acid is toxic to a wide range of plants and animals, particularly mice, chick embryos and chickens (Logrieco et al., 2003). Sorghum grains containing high levels of TA were associated with the human

62

haematological disorder known as onyalay (Bottalico and Logrieco, 1998). Less toxic compounds include alternariol (AOH) and alternariol monomethyl ether (AME), altenuene (AE) and altertoxin I, II and III (Logrieco et al., 2003). AOH and AME are usually found in combination and have some teratogenic and fetotoxic effects in pregnant mice (Bottalico and Logrieco, 1998). The production of one or more of these toxins has been reported in tomatoes (Bottalico and Logrieco, 1998), citrus fruits (Logrieco et al., 1990a), capsicum and melons (Bottalico and Logrieco, 2001), cereal grains (Logrieco et al., 1990b; Webley et al., 1997), rapeseed and olives (Logrieco et al., 2003), and apples (Robiglio and Lopez, 1995; Tournas and Stack, 2001). Maximum production of alternariol, its monomethyl ether and altenuene occurred at 258C and 0.98 aw (Magan et al., 1984). However, the optimum for tenuazonic acid production was reported as 0.90 aw at 258C (Etcheverry et al., 1994). Altertoxins I, II and III are produced by many Alternaria species, however, A. alternata is the most important producer. Altertoxins have received much attention due to their mutagenic activity, particularly that of altertoxin III, whose mutagenicity is approximately ten times lower than that of aflatoxin B1 (Bottalico and Logrieco, 1998). Ecology. Under the names Alternaria alternata and A. tenuis, this species has been reported from a very wide range of foods. However, the very broad species concept used by many authors is not accepted by Simmons (2007). The following reports undoubtedly include a number of other species. It is a major pathogen of fresh tomatoes (Harwig et al., 1979; Zitter and Wien, 1984; Pose et al., 2004), though other Alternaria species are probably involved here too. Spoilage of eggplants and peppers (Snowdon, 1991), apples (Combrink et al., 1985) and bananas stored under modified atmospheres (Wade et al., 1993) have been reported. Damage may occur also to a wide variety of fresh vegetables including beans, cauliflowers, cucumbers, melons, peas and potatoes (Webb and Mundt, 1978; Snowdon, 1991). A. alternata has been recorded frequently from a wide range of cereals, particularly wheat (Pitt et al., 1998b; Clear et al., 2005, Fakhrunnisa et al., 2006; Lugauskas et al., 2006 and references quoted in Pitt and Hocking, 1997), barley (Andersen et al., 1996; Fakhrunnisa et al., 2006; Medina et al., 2006 and references

5 Primary Keys and Miscellaneous Fungi

quoted in Pitt and Hocking, 1997), but also in rice, rapeseed, sunflower seed (Pozzi et al., 2005) and sorghum (Fakhrunnisa et al., 2006). In Australia and Southeast Asia, A. alternata has caused severe damage, spoilage and mycotoxin production in wheat and sorghum (Pitt et al., 1994; Webley et al., 1997 and our unpublished observations). This species has been recorded quite frequently from nuts, including peanuts, hazelnuts and pecans (see Pitt and Hocking, 1997). Other sources include soybeans, cold stored and frozen meat, biltong, bottled water and spices (Pitt and Hocking, 1997; Cabral and Fernadez Pinto, 2002). This species has occasionally been associated with gushing in beer (Niessen et al., 1992). Closely related species. Alternaria tenuissima (Kunze) Wiltshire differs from A. alternata by the formation of longer conidia (up to 60 mm) in unbranched chains (Simmons, 2007). The metabolite profiles of A. alternata and A. tenuissima are very similar. In a recent study, it was found that most A. tenuissima isolates examined produced alternariol, alternariol monomethyl ether and tenuzoic acid, whereas three of the A. alternata isolates produced altenuene, alternariol, alternariol monomethyl ether and altertoxin I (Andersen et al, 2002). Andersen and Frisvad (2004) reported A. tenuissima from mouldy tomatoes. In our experience this species is of common occurrence in cereals. References. Simmons (1967, 2007); Ellis (1971); Domsch et al. (1980).

Alternaria infectoria E.G. Simmons

Fig. 5.3

Colonies on CYA and MEA 60 mm diam or more, on CYA plane, low to moderately deep, mycelium dark grey to almost black, sometimes with a paler grey floccose overlay; reverse bluish black; on MEA moderately deep to deep, dense to floccose, of dark grey mycelium sometimes overlaid with lighter grey hyphae; reverse bluish black. On G25N, colonies 8–15 mm diam, of grey or greenish grey mycelium; reverse pinkish, grey or black. At 58C, at least germination, usually colonies of 2–5 mm diam formed. At 378C, colonies 5–25 mm diam, low to deep, grey, reverse dark. Colonies on DCMA 60 mm or more diam, usually with wide margins, low to subsurface,

5.4 Genus Alternaria

63

a

b

c

Fig. 5.3 Alternaria infectoria (a) colonies on PDA and DCMA, 7 d, 258C ; (b) conidia, bar ¼ 50 mm; (c) conidia, bar ¼ 20 mm

centrally usually sparse and floccose, aerial mycelium white to pale olive brown; reverse pale brown, brown or olive brown, sometimes darker in annular or irregular patches. Conidia on DCMA observed under the stereomicroscope characteristically formed in short, irregularly branched chains, forming clusters, with primary conidia borne from knobby hyphae, usually bearing secondary conidia borne from the tips, at the end of short (30–100 mm) hyphal lengths; conidia characteristically clavate (clubshaped) 20–50  7–14 mm, with up to 7 transverse septa and 1 or more longitudinal septa as well, or sometimes

irregularly septate, walls and septa thick and dark brown, and with walls sometimes conspicuously roughened. Distinctive features. After growth on DCMA, Alternaria infectoria is readily distinguished from A. alternata and other Alternaria species under the stereomicroscope. A. infectoria forms clusters of conidia borne in irregularly branching chains that, unlike those of A. alternata, have short hyphal segments between the conidia. Colonies on DCMA have low, wide margins, are less floccose than those of A. alternata and (apart from conidia) have paler colours.

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Taxonomy. This species was commonly known as the Alternaria state of Pleospora infectoria until Simmons (1986) provided a binomial name, A. infectoria. He showed that its teleomorph does not belong in Pleospora, as had been widely accepted, and erected a new genus, Lewia Barr and Simmons, for it (Simmons, 1986). Simmons (1986) reported that Alternaria species related to A. infectoria have very rarely been seen to produce a teleomorph; however, on several occasions we have seen Lewia teleomorphs in isolates of A. infectoria grown on autoclaved wheat grains. Physiology. No physiological studies on this species have been reported. Mycotoxins. Alternaria infectoria is chemically very different from other Alternaria species (Andersen et al., 2002). A. infectoria does not produce any of the known Alternaria metabolites (Andersen and Thrane, 1996) and shares only a few metabolites with other Alternaria species (Andersen et al., 2002). We have confirmed the absence of any toxin production by this species (unpublished). Ecology. Alternaria infectoria is most commonly associated with grains, having been reported from small grains in the United States (Bruce et al., 1984), Australian wheat (Pitt et al., 1998b), Danish barley (Andersen et al., 1996) and Norwegian grains (Kosiak et al., 2004). The lack of other reports of

5 Primary Keys and Miscellaneous Fungi

A. infectoria more likely reflects failure by most investigators to differentiate Alternaria species, rather than true rarity. It seems probable that this species may be confined to small grains. References. Simmons (1986, 2007).

5.5 Genus Arthrinium Kunze Arthrinium, in earlier literature more commonly known by its synonym Papularia, is not a common genus in foods, but has occasionally caused spoilage. Arthrinium produces relatively large, darkwalled (but not black) conidia borne solitarily, both terminally and laterally, on short, narrow conidiophores. Two species are treated here, A. phaeospermum and the Arthrinium state of Apiospora montagnei, distinguished from each other by conidial size.

Arthrinium phaeospermum (Corda) M.B. Ellis

Fig. 5.4

Papularia sphaerosperma (Pers.) Hohn. ¨

Colonies on CYA and MEA covering the whole Petri dish, mycelium low or floccose, coloured white or grey, sometimes with conspicuous areas of pink,

Fig. 5.4 Arthrinium phaeospermum (a) colony on CYA, 7 d, 258C; (b) conidia, bar ¼ 10 mm

5.6 Genus Aureobasidium

darkening in age; reverse yellow or brown. Colonies on G25N 10–18 mm diam, of white mycelium. Sometimes germination at 58C. No growth at 378C. Reproduction by solitary conidia, blown out from the ends of, or from denticles on the sides of, short, narrow, sometimes sinuous conidiophores, themselves borne in clusters from mother cells on natural substrates, but often singly in culture; conidia circular in plan view but elliptical from the side, 8–12  5–7 mm, dark brown, smooth walled, often with a narrow, hyaline band around the longest periphery. Distinctive features. The conidium of Arthrinium is distinctive: solitary, dark-walled, circular in plan but elliptical from the side, and often with a hyaline peripheral band. Cultivation on DCPA and under lights may assist sporulation. In the present context, Arthrinium is distinguished from Nigrospora by the latter’s production of jet black conidia entirely devoid of ornamentation. Physiology. Conidia of Arthrinium appear to be highly heat resistant. In apple juice, conidia survived a pasteurising process of 888C for 1.5 min, and in water, heating at 1058C for 2.5 min (Anon, 1967). No experimental details, i.e., numbers heated or come-up time, were reported, however, and a decimal reduction time cannot be calculated. Even so, survival of a heat treatment at 888C indicates that A. phaeospermum is uncommonly heat resistant for a conidial fungus. Optimum germination of A. phaeospermum occurs when the conidia are suspended in sterile saline solution at 208C for 15 min before plating onto 2% malt extract agar (Agut and Calvo, 2004). The authors also noted that the viability of Arthrinium conidia was low, making this genus relatively rare in the environment. Mycotoxins. Arthrinium species are not known to produce mycotoxins, however, reports of secondary metabolites with antimicrobial properties have been found for some Arthrinium species (quoted in Agut and Calvo, 2004). Ecology. Arthrinium phaeospermum (reported as Papularia sphaerosperma) has been recorded as a cause of spoilage of pasteurised apple juice (Anon., 1967). It has also been reported on barley, wheat flour, rice, pecans and airline meals (see Pitt and Hocking, 1997). It was isolated from Morelo cherries (Olszak, 1994) and Iranian barley (Asgari et al., 2004). A. phaeospermum has also been recorded as a pathogen

65

on sweet potatoes (Ravichandran and Sullia, 1983). We isolated A. phaeospermum at low levels from paddy rice, mung beans, soybeans, black beans and cashews in Thailand (Pitt et al., 1993). Closely related species. The Arthrinium state of Apiospora montagnei Sacc., also known as Papularia arundinis (Corda) Fr., is similar to A. phaeospermum in all characters examined, except for the production of smaller conidia, 6–8 mm long. Data sheets at the International Mycological Institute, Egham, Surrey, UK, record its isolation from white flour and molasses; it has also been reported from wheat (Pelhate, 1968), barley (Flannigan, 1969) and cashews (Pitt et al., 1993). References. Ellis (1971); Domsch et al. (1980).

5.6 Genus Aureobasidium Viala and G. Boyer Growth of Aureobasidium isolates is at first yeast-like, but, while remaining very low and mucoid, colonies spread rapidly and turn black in patches. Microscopically, hyphae as well as budding yeast-like cells are present; the latter are actually conidia. The conidia are borne from small denticles (minute projections) directly from the hyphal walls or from short lateral protrusions on the hyphae; characteristically 2–4 denticles on one cell will produce conidia synchronously. There are more than 10 species (Hermanides-Nijhof, 1977), but only one, Aureobasidium pullulans, is of importance in foods (Fig. 5.5).

Aureobasidium pullulans (de Bary) G. Arnaud

Fig. 5.5

Dematium pullulans de Bary Pullularia pullulans (de Bary) Berkhout

Colonies on CYA and MEA 25–35 mm diam, low and mucoid, faintly pink, becoming grey to black in areas at 7–10 days; reverse in similar colours. Colonies on G25N 10–12 mm diam, similar to those on CYA. At 58C, microcolonies to colonies up to 3 mm diam. No growth at 378C. Conidia borne on denticles directly from hyphae or sometimes small lateral protrusions; conidia yeast-like, primary ones borne from the denticles, usually measuring 10–16  3–6 mm, and secondary

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a

b

c

Fig. 5.5 Aureobasidium pullulans (a) colonies on CYA and MEA, 7 d, 258C; (b, c) conidia, bar ¼ 10 mm

ones by budding from the primaries, 7–10  3–5 mm, not adhering to each other, smooth walled. Distinctive features. Within the present context, Aureobasidium pullulans is readily recognised by its distinctive low, mucoid, white then pink to black colonies and yeast-like conidia. Several other genera have a similar appearance (Hermanides-Nijhof, 1977; Beh, 2007) but they rarely occur in foods. Taxonomy. Earlier literature discusses this species under the names Dematium pullulans and Pullularia pullulans. Mycotoxins. Mycotoxins are not known to be produced. Physiology. The temperature range for growth of Aureobasidium pullulans has been reported as 2–358C, with an optimum of 258C (Skou, 1969), but some earlier data, together with its abundance

in low temperature habitats, suggests that some strains grow down to –58C (Michener and Elliott, 1964). It is of common occurrence in salterns (Butinar et al., 2005a), so is likely to be a xerophile. Heat resistance is very low (Skou, 1969). The ability of A. pullulans to grow on sugars used in yeast taxonomy has been reported by de Hoog and Yurlova (1994) and Beh (2007). Ecology. A ubiquitous saprophyte from all sorts of moist and decaying environments, Aureobasidium pullulans has been reported from a very wide range of foods, but only rarely as a cause of spoilage. Its prevalence in frozen foods is noteworthy, being the predominant mould isolated from blueberry, apple and cherry pies by Kuehn and Gunderson (1963). It was the most frequently isolated mould in Canadian icewine must, but played no role in fermentating

5.7 Genus Bipolaris

must (Subden et al., 2003). The surface of healthy grapes is predominately colonised by A. pullulans (Fleet, 2003a; Prakichaiwattana et al., 2004; Beh, 2007). A. pullulans has been reported in yoghurt in a survey of a Slovakian dairy plant (Pieckova et al., 2002) and has been associated with the spoilage of cold stored meat and cheese (see Pitt and Hocking, 1997). A. pullulans has been recorded in a variety of fresh foods and commodities including fresh vegetables, cabbage, strawberries, grapes, citrus and pasteurised orange juice (see Pitt and Hocking, 1997). Other records include shrimp, green olives, barley, wheat and flour, oats and nuts (see Pitt and Hocking, 1997). References. Hermanides-Nijhof (1977); Domsch et al. (1980); de Hoog and Yurlova (1994).

5.7 Genus Bipolaris Shoemaker In separating unrelated species from Helminthosporium, Shoemaker (1959) revised the genus Drechslera S. Ito and erected Bipolaris. The fact that species of Bipolaris (in the sense of Shoemaker, 1959) were associated with two distinct teleomorph types, and that correlated with morphological differences, led to segregation of Exserohilum from Bipolaris by Leonard and Suggs (1974). These genera produce long, large conidia with transverse septa only; the septa are thick and quite different from those in e.g. Trichoconiella. In Drechslera, conidia are cylindrical and germinate at any cell, while conidia of Bipolaris and Exserohilum are fusoid, i.e. gradually narrowing, and conidia germinate only from the end cells (bipolar germination). In Bipolaris, germ tubes develop roughly along the line of the conidium axis, while in Exserohilum, the germ tube is often offset from the axis and develops at an angle (Alcorn, 1983). Exserohilum species are apparently rare in foods, although E. rostratum (Drechsler) K.J. Leonard and Suggs has been reported from sorghum (Usha et al., 1994). The literature on these genera is voluminous and complex. The most useful publications are by Alcorn (1983) and Sivanesan (1987). One common species of Bipolaris, B. maydis, is treated here (Fig. 5.6).

67

Bipolaris maydis (Nishik. and C. Miyake) Shoemaker Fig. 5.6 Colonies on CYA and MEA 45–55 mm diam, plane, deeply floccose, of light to dark grey mycelium; reverse dark brown to dark blue black. On G25N, colonies 2–6 mm diam. At 58C, germination. At 378C, colonies 5–15 mm diam, usually low, dense and wrinkled, of grey mycelium, reverse deep blue black. Colonies on DCMA 30–40 mm diam, plane, of sparse to dense, floccose grey mycelium, reverse brown, grey or black. Conidia borne singly from nodes on knobby or geniculate mid to dark brown hyphae, usually slightly curved, (40–)50–80  12–16 mm, with six to nine inconspicuous septa and with smooth, dark, thick walls. Distinctive features. Large conidia, often slightly curved, of fusoid shape and with several thick transverse septa are characteristic of this genus. Conidial curvature and shape distinguish it from Drechslera, while conidial length and the presence of more than three, thick septa allow distinction from Curvularia. The species treated here is common on maize. Taxonomy. Some species of Bipolaris produce teleomorphs, classified in the genus Cochliobolus. Physiology. Species of Bipolaris sporulate significantly better when grown on substrates containing cellulose (Pratt, 2006). Mycotoxins. No significant mycotoxins are known. Ecology. We have isolated Bipolaris maydis from 1% of Thai maize kernels, and less frequently from Thai sorghum and mung bean samples (Pitt et al., 1993, 1994). It was also found at similar levels in commodities from Indonesia and the Philippines (Pitt et al., 1998a). Other records of Bipolaris in foods are rare. However, references to Drechslera species in foods often have referred to taxa correctly classified in Bipolaris, e.g. in rice, B. oryzae (Breda de Haan) Shoemaker; in barley, B. australiensis (M.B. Ellis) Tsuda and Uemaya, B. maydis and B. setariae (Sawada) Shoemaker; in peanuts, B. spicifera (Bainier) Subram.; in coriander seed, B. bicolor (Mitra) Shoemaker; and in avocados, B. setariae (see Pitt and Hocking, 1997). References. Alcorn (1983); Sivanesan (1987).

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a

b

c

Fig. 5.6 Bipolaris maydis (a) colonies on MEA and DCMA, 7 d, 258C; (b, c) conidiophores and conidia, bars ¼ 10 mm

5.8 Genus Botrytis P. Micheli: Fr. Botrytis is a common genus in the temperate zones, where it occurs mainly as a pathogen on a variety of plant crops. Vegetables and small berry fruits are particularly susceptible. Invasion may occur before maturity or postharvest, both in transport and in storage. Onions and other Allium species and grapes are the most susceptible crops. In the latter, it is notable that the disease is sometimes encouraged. Grapes affected by Botrytis, in this circumstance called ‘‘the noble rot’’, are used in the production of certain high quality sweet wines in France, Germany, Australia and other countries. Botrytis species are characterised by the production of conidia on pegs from spherical swellings. The most commonly encountered species in foods

is known as Botrytis cinerea, although it is probable that this name includes a group of related species rather than a single well defined taxon (Coley-Smith et al., 1980).

Botrytis cinerea Pers.

Fig. 5.7

Colonies on CYA and MEA covering the whole Petri dish, floccose, growth sometimes patchy or irregular, mycelium white, becoming grey to dark grey as conidiogenesis proceeds; reverse pale to grey. On G25N, colonies 10–18 mm diam, irregular in outline, floccose centrally or in patches, becoming grey; reverse grey. At 58C, colonies up to 5 mm diam produced, low and sparse. No growth at 378C.

5.8 Genus Botrytis

69

Fig. 5.7 Botrytis cinerea (a) colony on CYA, 7 d, 258C; (b) conidiophores and conidia, bar = 25 mm

Conidiophores borne from aerial hyphae, stipes of indeterminate length, each bearing terminally an irregular cluster of short branches, 10–30 mm long, with swollen spherical apices, 8–10 mm diam; conidia borne singly from these apices on denticles (small pegs), ellipsoidal, 8–12 mm long, smooth walled, not released at maturity. Distinctive features. Solitary conidia borne on denticles from terminally swollen, short branches are characteristic of Botrytis. Physiology. Growth has been reported at 0.93 aw (Snow, 1949) and at 0.90 aw (Jarvis, 1977). Lahlali et al. (2007) observed no growth of Botrytis cinerea at 0.93 aw in the presence of NaCl or at 0.89 aw in the presence of nonionic solute. Reported growth temperatures are rather variable, with minima from –2 to 58C or even 128C, maxima 28–358C, and optima of 22–258C (Domsch et al., 1980). B. cinerea will grow from pH 2–8 (Jarvis, 1977) and in O2 concentrations down to 1% (Follstad, 1966). The effect of low temperatures and controlled atmosphere storage on the growth of B. cinerea on susceptible fruits and vegetables has been reported (Reyes, 1990). B. cinerea produces pectinases (polygalacturonase) and the cell wall degrading enzymes cellulase and xylanase (El-Habbaa, 2003). Mycotoxins. Mycotoxin production has not been reported – fortunately, in view of this species role in speciality wine production.

Ecology. Botrytis cinerea is a virulent pathogen, the cause of rots in many kinds of fresh fruits (Snowdon, 1990). It is the most important cause of disease in grapes, both before harvest and in storage and the most common cause of spoilage in berries such as strawberries, blueberries, raspberries and blackberries (Snowdon, 1990; Tournas and Katsoudas, 2005), and also causes large losses in apples and pears, tomatoes, stone fruits and kiwi fruit (see Pitt and Hocking, 1997). A gel-like turbidity can be caused in raspberry juice, making filtration difficult (Will et al., 1992). For a review of many aspects of occurrence, pathogenicity and control, see Elad et al. (2004). Although vegetable spoilage is perhaps less important, the range of crops affected is very large, and includes asparagus, beans, cabbages, carrots, celery, melons and potatoes (Snowdon, 1991; Lugauskas et al., 2005; Tournas, 2005). It has also been isolated from a wide variety of dried or processed foods, but here its role is uncertain and it is probably present only as a contaminant. Other important species. Several other species of Botrytis are commonly occurring pathogens. Of interest here are B. allii Munn, B. byssoides Walker and the Botrytis state of Sclerotinia porri van Beyma, all of which cause spoilage of onions and leeks (Ellis, 1971; Snowdon, 1990). Botrytis allii grows optimally at 20–258C, with a minimum below 58C and a maximum near 358C. The optimum aw for growth is in excess of 0.99. Growth

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occurs down to 0.96 aw in media containing NaCl and 0.93 aw in media containing KCl or sucrose, or on onion leaves (Alderman and Lacy, 1984). References. Ellis (1971); Coley-Smith et al. (1980); Domsch et al. (1980); Elad et al. (2004).

5.9 Genus Chaetomium Kunze Chaetomium is the only genus of Ascomycetes which produces black ascocarps and which is commonly encountered in foods. The ascocarps in Chaetomium are perithecia, similar in appearance to cleistothecia, but having an opening (ostiole) through which the ascospores are released. There is a superficial resemblance to the genus Phoma, which produces black

conidiomata (pycnidia); ascospores in Chaetomium are often freed from the ascus before being discharged singly from the perithecium, and hence could be mistaken for conidia. Chaetomium is readily distinguished from Phoma by the presence of stout black or dark coloured hyphae attached to the perithecial walls, which are visible in culture under the low power microscope. Chaetomium species are notable as producers of cellulases and so commonly occur on wood and paper products. They are relatively common in some kinds of foods. C. globosum is the most frequently encountered species and is described here as representative of the genus. In addition descriptions of C. brasiliense and C. funicola, both quite common in tropical commodities, are provided here.

Key to Chaetomium species included here 1

Colonies on CYA at 378C much larger than at 258C Colonies on CYA at 378C smaller than at 258C or absent

2 (1)

Colonies on CYA and MEA at 258C less than 50 mm diam, ascospores 5–7 mm long Colonies on CYA and MEA at 258C usually covering the whole Petri dish, ascospores 8–10 mm long

Chaetomium brasiliense Bat. & Pontual

Fig. 5.8

Colonies on CYA and MEA 25–35 mm diam, on CYA low to moderately deep and dense, of white to grey mycelium; reverse sometimes pale, more commonly dark grey to almost black; on MEA low and sparse, dark grey, with some paler grey aerial hyphae; reverse dark grey. No growth on G25N or germination at 58C. Colonies at 378C covering the whole Petri dish, 65 mm or more diam, general appearance similar to those on CYA at 258C; reverse very dark grey to greyish black (Fig. 5.8). Reproductive structures perithecia not observed on CYA or MEA at 258C, but on CYA at 378C abundant, forming a deep, contiguous layer on the agar surface, most readily observed at colony margins, especially between colonies, surrounded by radiating long, stout hyphae, straight at first but at maturity curling into long corkscrew shapes, 75–100 mm diam, maturing in 7 days at colony centres; asci not seen, ascospores

C. brasiliense 2 C. funicola C. globosum

subspheroidal to broadly ellipsoidal, 6–8 mm long, brown, with smooth walls. Conidia not produced. Distinctive features. Unlike the other Chaetomium species considered here, C. brasiliense grows much more rapidly at 378C than 258C, and perithecia are formed in much greater numbers at the higher temperature. Perithecia are black, but under the stereomicroscope appear surrounded by grey hyphae. Ascospores are smaller than those of C. globosum. Physiology. No studies on physiology are known to us. However, the description above indicates that this species has a high temperature optimum for growth. Mycotoxins. Significant mycotoxins are not known to be produced. Ecology. We have isolated this species not infrequently from tropical sources. From Thailand, 1% of mung beans, black beans, paddy rice grains and cashew kernels, and 7–10% of soy bean and sorghum samples, but <1% of individual seeds, contained C. brasiliense (Pitt et al., 1993, 1994). Similar figures were obtained from Indonesian commodities (Pitt et al., 1998a). In samples from the Philippines, this species was found in maize, peanuts, soybeans, mung beans,

5.9 Genus Chaetomium

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Fig. 5.8 Chaetomium brasiliense (a) colony on CYA, 7 d, 258C; (b) perithecium in situ, bar ¼ 25 mm; (c) asci, bar ¼ 10 mm; (d) ascospores, bar ¼ 5 mm

black pepper, paddy and milled rice, all near 1% of total particles examined (our unpublished data). Reference. Von Arx et al. (1986).

Chaetomium funicola Cooke

Fig. 5.9

Colonies on CYA 20–25 mm diam, plane, of low and sparse white mycelium, sometimes pale grey centrally, enveloping abundant developing perithecia, reverse pale or greyish brown. Colonies on MEA 35–40 mm diam, plane, sparse, low to somewhat floccose, mycelium white or grey, enveloping developing perithecia; reverse pale to greyish olive or pale brown. On G25N, usually no growth. At 58C, no germination. At 378C, usually no growth.

Perithecia borne in a layer on the agar surface, 120–160 mm diam, surrounded by almost straight, dark walled, rough to spiky hyphae; asci maturing in 14 days, breaking down immediately, ascospores ellipsoidal, 5–7  3–5 mm, grey or brown, sometimes with one or two internal oil droplets visible, often collapsing in age, smooth walled. Distinctive features. Chaetomium funicola grows only slowly at 258C and usually not at 378C. Ascospores are smaller than those of C. globosum. Physiology. No studies are known to us. Domsch et al. (1980) indicated that perithecial production is maximal at about 208C, indicating that this species is better adapted to low than high temperatures.

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Fig. 5.9 Chaetomium funicola (a) colonies on CYA and MEA, 7 d, 258C; (b) perithecium in situ, bar ¼ 50 mm; (c) ascospores, bar ¼ 5 mm

Mycotoxins. Mycotoxin production has not been reported. Ecology. Despite its low temperature optimum, we repeatedly isolated Chaetomium funicola from tropical commodities. It was present in 23% of Indonesian soybean samples, ranging up to 50% infection in some samples, and in 3% of beans overall (our unpublished data). Lower levels of infection (1% or more overall) were observed in peanuts, cashews, copra, mung beans and sorghum from Thailand (Pitt et al., 1993, 1994), in mung beans and maize from Indonesia, and paddy rice from the Philippines (Pitt et al., 1998a and our unpublished data). References. Domsch et al. (1980); von Arx et al. (1986).

Chaetomium globosum Kunze

Fig. 5.10

Colonies on CYA and MEA covering the whole Petri dish, on CYA low and sparse, of scanty white mycelium and conspicuous though usually sparse black perithecia, ca. 0.2 mm diam; on MEA growth more dense but still low, coloured grey or greenish black from hyphae enveloping abundant perithecia; reverse on both media usually brown. On G25N, colonies less than 5 mm diam produced. No growth at 58C. At 378C, colonies usually 20–30 mm diam, of white mycelium. Reproductive structures perithecia, black, 150–200 mm diam, with numerous stout, dark hyphae appended; ascospores produced after 1–2 weeks,

5.10 Genus Chrysonilia

73

Fig. 5.10 Chaetomium globosum (a) colony on CYA, 7 d, 258C; (b) perithecium in situ, bar ¼ 100 mm; (c) ascospores, bar ¼ 5 mm

spheroidal, broadly ellipsoidal or apiculate, commonly 8–10 mm long, smooth walled. Conidia not generally produced. Distinctive features. In contrast to Chaetomium brasiliense, C. globosum grows much more rapidly at 258C than 378C. Ascospores are larger than those of C. funicola. Physiology. Chapman and Fergus (1975) reported that Chaetomium globosum ascospores germinated from a minimum temperature of 4–108C to a maximum of 388C, and most rapidly at 24–388C. They also reported germination over the whole pH range tested, 3.5–7. Heat resistance was low: 1% of ascospores survived 10 min at 558C, but none survived at 578C for a similar period. Ahammed et al. (2005) observed maximum sporulation at 288C and pH 5.0 on a medium containing sucrose and thiamine. C. globosum can grow down to 0.94 aw in soil (Kouyeas, 1964). Mycotoxins. Chaetomium globosum produces chaetoglobsins A and C, moderately toxic compounds reported to be produced in building materials (Fogle et al., 2007). Their relevance to foods is unknown. Ecology. This species has been isolated from a variety of commodities, particularly wheat, barley, rice, beans and soybeans. Chaetomium globosum has also been recorded in nuts (see Pitt and Hocking, 1997) and spices (Mandeel, 2005). We isolated it

frequently from tropical commodities: from 12 to 17% of maize samples, 14 to 32% of mung bean samples and 33 to 35% of soybean samples from Thailand, Indonesia and the Philippines, with overall infection rates of 1–4% of all particles examined. Relatively high infection levels were also encountered in Thai cashews, copra and paddy rice. Kemiri (candle) nuts from Indonesia were highly infected: 37% of samples, up to 40% in infected samples and 9% of nuts overall (Pitt et al., 1998a). This species has not been reported to cause food spoilage, although it has recently been implicated in causing disease in pears in Egypt (Ismail and Abdalla, 2005). References. Domsch et al. (1980); von Arx et al. (1986).

5.10 Genus Chrysonilia Arx Chrysonilia was erected by von Arx (1981a) to accommodate Monilia sitophila, for many years known as ‘‘the red bread mould’’. Chrysonilia sitophila is characterised by very rapid growth and the production of conidia in chains, cut off from the apices of undifferentiated hyphae. This species is readily recognisable in Petri dish culture: sparse, white to pink mycelium spreads rapidly across the dish and up the walls, then within 3 days sheds

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Fig. 5.11 Chrysonilia sitophila (a) colony on CYA, 7 d, 258C; (b) conidia, bar ¼ 10 mm

enormous numbers of orange conidia outside the dish. This attribute makes it a particularly troublesome laboratory contaminant. Cultivation in Petri dishes is therefore not recommended: observation of these characters in a culture tube is sufficient for positive identification. The teleomorph of this species is Neurospora sitophila, a well-known Ascomycete which has been of great value in genetic studies. It is not found in foods, however.

Chrysonilia sitophila (Mont.) Arx

Fig. 5.11

Monilia sitophila (Mont.) Sacc.

Teleomorph: Neurospora sitophila Shear and B.O. Dodge Colonies in 28 ml McCartney bottles or test tubes distinguished by pale pink floccose growth, filling the entire bottle, then turning salmon, first at the bottle neck or tube plug and subsequently throughout, as sporulation occurs. On CYA and MEA, colonies covering the whole Petri dish, mycelium pink, reaching the lid in tufts or patches and all around the rim, producing vast numbers of salmon conidia at or near the rim and sometimes for several millimetres beyond it, and shedding them profusely; reverse salmon or pink. On G25N, colonies up to 30 mm diam produced,

low, dense and mucoid. No growth at 58C. At 378C, colonies covering the whole Petri dish, similar to at 258C. Reproductive structures arthroconidia, cut off in succession from the apices of branching hyphae; conidia at maturity variable in size and shape, spherical to ellipsoidal, pale orange, 6–15 mm diam, with thin smooth walls. Distinctive features. See remarks above. Physiology. According to Panasenko (1967), Chrysonilia sitophila is capable of growth down to 0.88–0.90 aw. Mycotoxins. No mycotoxins have been recorded for this species. Ecology. For long a common sight in bakeries and on bread, Chrysonilia sitophila is less commonly encountered now, but can still be a great source of trouble as a p